Male arthropod killing factors and methods of use thereof
The present disclosure relates to genetically modified arthropods, genetically modified bacteria, and methods for controlling and/or reducing arthropod populations.
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This application is a 371 of PCT International Application No. PCT/US2019/025936, filed Apr. 5, 2019, which claims the benefit of U.S. Provisional Patent Application Ser. No. 62/652,982, filed Apr. 5, 2018, which are expressly incorporated herein by reference.
STATEMENT REGARDING FEDERALLY SPONSORED RESEARCHThis invention was made with government support under Grant Nos. HD086833, AI133522, AI132581, CA068485, DK020593, DK058404, DK059637 and EY008126 awarded by the National Institutes of Health and under Grant No. 1456778 awarded by the National Science Foundation. The government has certain rights in the invention.
FIELDThe present disclosure relates to genetically modified arthropods, genetically modified bacteria, and methods for controlling and/or reducing arthropod populations.
BACKGROUNDWolbachia are maternally-transmitted bacteria that infect almost half of all arthropod species and many nematode species. In arthropods, these bacteria often selfishly manipulate host reproduction to enhance the fitness of infected females, thereby facilitating their own transmission and spread through the host population. Despite significant impacts of Wolbachia on animal reproduction, evolution, and vector control, the bacterial genes underlying most of these reproductive manipulations remain elusive. One such phenotype is male killing, where the sons of infected females are selectively killed. What is needed are novel compositions and methods that can recapitulate the male killing phenotype in order to improve vector control and thus reduce transmission of vector-borne diseases.
The compounds, compositions, and methods disclosed herein address these and other needs.
SUMMARYDisclosed herein are genetically modified arthropods and genetically modified bacteria useful for controlling and/or reducing populations of arthropods (for example, insects). The inventors have identified a gene, hereafter denoted WO male killing (wmk) [or WD0626 (also referred to as locus number WD_RS02815)], that causes male lethality when transgenically expressed, which leads to a female-biased sex ratio. These male arthropod-killing genes encoding male arthropod killing factors are used to genetically modify arthropods in order to reduce a population of target arthropods, such as vectors of disease or agricultural pests. In addition, the methods of using these male arthropod-killing factors can be combined with additional methods for vector control, for example, sterile insect techniques (SIT) or incompatible insect techniques (IIT).
In some aspects, disclosed herein is a genetically modified arthropod, said arthropod comprising:
-
- a gene encoding a male arthropod killing factor; and
- a promoter operably linked to the gene encoding the male arthropod killing factor;
- wherein the expression of the gene encoding the male arthropod killing factor in arthropod embryos causes a reduction in viable surviving male offspring in comparison to arthropod embryos not expressing the gene encoding the male arthropod killing factor.
In some embodiments, the gene encoding the male arthropod killing factor is from a bacterium. In some embodiments, the gene encoding the male arthropod killing factor is from a phage or prophage. In some embodiments, the gene encoding the male arthropod killing factor is from Wolbachia (or prophage WO or phage WO). In some embodiments, the male arthropod killing factor is wmk (WD0626) or its homologs. In some embodiments, the male arthropod killing factor comprises the amino acid sequence SEQ ID NO:1, or a variant thereof.
In some embodiments, the reduction in viable male offspring is greater than 10%.
In some embodiments, the arthropod is an insect. In some embodiments, the insect is selected from the genera consisting of Aedes, Culex and Anopheles. In some embodiments, the insect is selected from the group consisting of Aedes albopictus, Aedes aegypti and Aedes polynesiensis. In some embodiments, the insect is Drosophila suzukii.
In some aspects, disclosed herein is a method for controlling a population of target arthropods, comprising:
-
- providing a gene encoding a male arthropod killing factor, and a promoter operably linked to the gene encoding the male arthropod killing factor;
- transforming a population of arthropods with the gene encoding the male arthropod killing factor and the promoter operably linked to the gene encoding the male arthropod-killing factor; and
- releasing the population of arthropods amongst a population of target arthropods, wherein the release of the arthropods reduces the population of target arthropods.
In some aspects, disclosed herein is a method for controlling a population of target arthropods, comprising:
-
- providing a gene encoding a male arthropod killing factor, and a promoter operably linked to the gene encoding the male arthropod killing factor;
- genetically transforming a bacteria, phage, or prophage with the gene encoding the male arthropod killing factor operably linked to the promoter;
- transinfecting a population of arthropods with the bacteria, phage, or prophage; and
- releasing the population of arthropods amongst a population of target arthropods, wherein the release of the arthropods reduces the population of target arthropods.
In some embodiments, the method further comprises providing an additional method of arthropod control. In some embodiments, the additional method of arthropod control is a sterile insect technique (SIT). In some embodiments, the additional method of arthropod control is an incompatible insect technique (IIT).
The accompanying figures, which are incorporated in and constitute a part of this specification, illustrate several aspects described below.
Disclosed herein are genetically modified arthropods and genetically modified bacteria useful for controlling and/or reducing populations of arthropods (for example, insects). Further disclosed herein are genetically modified bacteria, genetically modified phage, and/or genetically modified prophage for controlling and/or reducing populations of arthropods (for example, insects). Also disclosed herein is an arthropod transinfected with a male-killing microbe, phage or bacteria that naturally infects another host. The inventors have identified a gene, hereafter denoted WO male killing (wmk) (or locus WD0626 in the wMel Wolbachia genome), that causes male lethality when transgenically expressed, which leads to a female-biased sex ratio. These male arthropod-killing genes encoding male arthropod killing factors are used to genetically modify arthropods in order to reduce a population of target arthropods. In addition, the methods of using these male arthropod killing factors can be combined with additional methods for vector control, such as sterile insect techniques or incompatible insect techniques.
Reference will now be made in detail to the embodiments of the invention, examples of which are illustrated in the drawings and the examples. This invention may, however, be embodied in many different forms and should not be construed as limited to the embodiments set forth herein.
Unless defined otherwise, all technical and scientific terms used herein have the same meaning as commonly understood to one of ordinary skill in the art to which this disclosure belongs. The term “comprising” and variations thereof as used herein is used synonymously with the term “including” and variations thereof and are open, non-limiting terms. Although the terms “comprising” and “including” have been used herein to describe various embodiments, the terms “consisting essentially of” and “consisting of” can be used in place of “comprising” and “including” to provide for more specific embodiments and are also disclosed. As used in this disclosure and in the appended claims, the singular forms “a”, “an”, “the”, include plural referents unless the context clearly dictates otherwise.
The following definitions are provided for the full understanding of terms used in this specification.
Terminology
The term “nucleic acid” as used herein means a polymer composed of nucleotides, e.g. deoxyribonucleotides or ribonucleotides.
The terms “ribonucleic acid” and “RNA” as used herein mean a polymer composed of ribonucleotides.
The terms “deoxyribonucleic acid” and “DNA” as used herein mean a polymer composed of deoxyribonucleotides.
The term “oligonucleotide” denotes single- or double-stranded nucleotide multimers of from about 2 to up to about 100 nucleotides in length. Suitable oligonucleotides may be prepared by the phosphoramidite method described by Beaucage and Carruthers, Tetrahedron Lett., 22:1859-1862 (1981), or by the triester method according to Matteucci, et al., J. Am. Chem. Soc., 103:3185 (1981), both incorporated herein by reference, or by other chemical methods using either a commercial automated oligonucleotide synthesizer or VLSIPS™ technology. When oligonucleotides are referred to as “double-stranded,” it is understood by those of skill in the art that a pair of oligonucleotides exist in a hydrogen-bonded, helical array typically associated with, for example, DNA. In addition to the 100% complementary form of double-stranded oligonucleotides, the term “double-stranded,” as used herein is also meant to refer to those forms which include such structural features as bulges and loops, described more fully in such biochemistry texts as Stryer, Biochemistry, Third Ed., (1988), incorporated herein by reference for all purposes.
The term “polynucleotide” refers to a single or double stranded polymer composed of nucleotide monomers. In some embodiments, the polynucleotide is composed of nucleotide monomers of generally greater than 100 nucleotides in length and up to about 8,000 or more nucleotides in length.
The term “polypeptide” refers to a compound made up of a single chain of D- or L-amino acids or a mixture of D- and L-amino acids joined by peptide bonds.
The term “complementary” refers to the topological compatibility or matching together of interacting surfaces of a probe molecule and its target. Thus, the target and its probe can be described as complementary, and furthermore, the contact surface characteristics are complementary to each other.
The term “hybridization” refers to a process of establishing a non-covalent, sequence-specific interaction between two or more complementary strands of nucleic acids into a single hybrid, which in the case of two strands is referred to as a duplex.
The term “anneal” refers to the process by which a single-stranded nucleic acid sequence pairs by hydrogen bonds to a complementary sequence, forming a double-stranded nucleic acid sequence, including the reformation (renaturation) of complementary strands that were separated by heat (thermally denatured).
The term “melting” refers to the denaturation of a double-stranded nucleic acid sequence due to high temperatures, resulting in the separation of the double strand into two single strands by breaking the hydrogen bonds between the strands.
The term “promoter” or “regulatory element” refers to a region or sequence determinants located upstream or downstream from the start of transcription and which are involved in recognition and binding of RNA polymerase and other proteins to initiate transcription. Promoters need not be of bacterial origin, for example, promoters derived from viruses or from other organisms can be used in the compositions, systems, or methods described herein.
A polynucleotide sequence is “heterologous” to a second polynucleotide sequence if it originates from a foreign species, or, if from the same species, is modified by human action from its original form. For example, a promoter operably linked to a heterologous coding sequence refers to a coding sequence from a species different from that from which the promoter was derived, or, if from the same species, a coding sequence which is different from naturally occurring allelic variants.
The term “recombinant” refers to a human manipulated nucleic acid (e.g. polynucleotide) or a copy or complement of a human manipulated nucleic acid (e.g. polynucleotide), or if in reference to a protein (i.e, a “recombinant protein”), a protein encoded by a recombinant nucleic acid (e.g. polynucleotide). In embodiments, a recombinant expression cassette comprising a promoter operably linked to a second nucleic acid (e.g. polynucleotide) may include a promoter that is heterologous to the second nucleic acid (e.g. polynucleotide) as the result of human manipulation (e.g., by methods described in Sambrook et al., Molecular Cloning—A Laboratory Manual, Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y., (1989) or Current Protocols in Molecular Biology Volumes 1-3, John Wiley & Sons, Inc. (1994-1998)). In another example, a recombinant expression cassette may comprise nucleic acids (e.g. polynucleotides) combined in such a way that the nucleic acids (e.g. polynucleotides) are extremely unlikely to be found in nature. For instance, human manipulated restriction sites or plasmid vector sequences may flank or separate the promoter from the second nucleic acid (e.g. polynucleotide). One of skill will recognize that nucleic acids (e.g. polynucleotides) can be manipulated in many ways and are not limited to the examples above.
The term “expression cassette” refers to a nucleic acid construct, which when introduced into a host cell, results in transcription and/or translation of a RNA or polypeptide, respectively. In embodiments, an expression cassette comprising a promoter operably linked to a second nucleic acid (e.g. polynucleotide) may include a promoter that is heterologous to the second nucleic acid (e.g. polynucleotide) as the result of human manipulation (e.g., by methods described in Sambrook et al., Molecular Cloning—A Laboratory Manual, Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y., (1989) or Current Protocols in Molecular Biology Volumes 1-3, John Wiley & Sons, Inc. (1994-1998)). In some embodiments, an expression cassette comprising a terminator (or termination sequence) operably linked to a second nucleic acid (e.g. polynucleotide) may include a terminator that is heterologous to the second nucleic acid (e.g. polynucleotide) as the result of human manipulation. In some embodiments, the expression cassette comprises a promoter operably linked to a second nucleic acid (e.g. polynucleotide) and a terminator operably linked to the second nucleic acid (e.g. polynucleotide) as the result of human manipulation. In some embodiments, the expression cassette comprises an endogenous promoter. In some embodiments, the expression cassette comprises an endogenous terminator. In some embodiments, the expression cassette comprises a synthetic (or non-natural) promoter. In some embodiments, the expression cassette comprises a synthetic (or non-natural) terminator.
The terms “identical” or percent “identity,” in the context of two or more nucleic acids or polypeptide sequences, refer to two or more sequences or subsequences that are the same or have a specified percentage of amino acid residues or nucleotides that are the same (i.e., about 60% identity, preferably 61%, 62%, 63%, 64%, 65%, 66%, 67%, 68%, 69%, 70%, 71%, 72%, 73%, 74%, 75%, 76%, 77%, 78%, 79%, 80%, 81%, 82%, 83%, 84%, 85%, 86%, 87%, 88%, 89%, 90%, 91%, 92%, 93%, 94%, 95%, 96%, 97%, 98%, 99% or higher identity over a specified region when compared and aligned for maximum correspondence over a comparison window or designated region) as measured using a BLAST or BLAST 2.0 sequence comparison algorithms with default parameters described below, or by manual alignment and visual inspection (see, e.g., NCBI web site or the like). Such sequences are then said to be “substantially identical.” This definition also refers to, or may be applied to, the compliment of a test sequence. The definition also includes sequences that have deletions and/or additions, as well as those that have substitutions. As described below, the preferred algorithms can account for gaps and the like. Preferably, identity exists over a region that is at least about 10 amino acids or 20 nucleotides in length, or more preferably over a region that is 10-50 amino acids or 20-50 nucleotides in length. As used herein, percent (%) amino acid sequence identity is defined as the percentage of amino acids in a candidate sequence that are identical to the amino acids in a reference sequence, after aligning the sequences and introducing gaps, if necessary, to achieve the maximum percent sequence identity. Alignment for purposes of determining percent sequence identity can be achieved in various ways that are within the skill in the art, for instance, using publicly available computer software such as BLAST, BLAST-2, ALIGN, ALIGN-2 or Megalign (DNASTAR) software. Appropriate parameters for measuring alignment, including any algorithms needed to achieve maximal alignment over the full-length of the sequences being compared can be determined by known methods.
For sequence comparisons, typically one sequence acts as a reference sequence, to which test sequences are compared. When using a sequence comparison algorithm, test and reference sequences are entered into a computer, subsequence coordinates are designated, if necessary, and sequence algorithm program parameters are designated. Preferably, default program parameters can be used, or alternative parameters can be designated. The sequence comparison algorithm then calculates the percent sequence identities for the test sequences relative to the reference sequence, based on the program parameters.
One example of algorithm that is suitable for determining percent sequence identity and sequence similarity are the BLAST and BLAST 2.0 algorithms, which are described in Altschul et al. (1977) Nuc. Acids Res. 25:3389-3402, and Altschul et al. (1990) J. Mol. Biol. 215:403-410, respectively. Software for performing BLAST analyses is publicly available through the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov/). This algorithm involves first identifying high scoring sequence pairs (HSPs) by identifying short words of length W in the query sequence, which either match or satisfy some positive-valued threshold score T when aligned with a word of the same length in a database sequence. T is referred to as the neighborhood word score threshold (Altschul et al. (1990) J. Mol. Biol. 215:403-410). These initial neighborhood word hits act as seeds for initiating searches to find longer HSPs containing them. The word hits are extended in both directions along each sequence for as far as the cumulative alignment score can be increased. Cumulative scores are calculated using, for nucleotide sequences, the parameters M (reward score for a pair of matching residues; always >0) and N (penalty score for mismatching residues; always <0). For amino acid sequences, a scoring matrix is used to calculate the cumulative score. Extension of the word hits in each direction are halted when: the cumulative alignment score falls off by the quantity X from its maximum achieved value; the cumulative score goes to zero or below, due to the accumulation of one or more negative-scoring residue alignments; or the end of either sequence is reached. The BLAST algorithm parameters W, T, and X determine the sensitivity and speed of the alignment. The BLASTN program (for nucleotide sequences) uses as defaults a wordlength (W) of 11, an expectation (E) or 10, M=5, N=−4 and a comparison of both strands. For amino acid sequences, the BLASTP program uses as defaults a wordlength of 3, and expectation (E) of 10, and the BLOSUM62 scoring matrix (see Henikoff and Henikoff (1989) Proc. Natl. Acad. Sci. USA 89:10915) alignments (B) of 50, expectation (E) of 10, M=5, N=−4, and a comparison of both strands.
The BLAST algorithm also performs a statistical analysis of the similarity between two sequences (see, e.g., Karlin and Altschul (1993) Proc. Natl. Acad. Sci. USA 90:5873-5787). One measure of similarity provided by the BLAST algorithm is the smallest sum probability (P(N)), which provides an indication of the probability by which a match between two nucleotide or amino acid sequences would occur by chance. For example, a nucleic acid is considered similar to a reference sequence if the smallest sum probability in a comparison of the test nucleic acid to the reference nucleic acid is less than about 0.2, more preferably less than about 0.01.
The phrase “codon optimized” as it refers to genes or coding regions of nucleic acid molecules for the transformation of various hosts, refers to the alteration of codons in the gene or coding regions of polynucleic acid molecules to reflect the typical codon usage of a selected organism without altering the polypeptide encoded by the DNA. Such optimization includes replacing at least one, or more than one, or a significant number, of codons with one or more codons that are more frequently used in the genes of that selected organism. For example, the sequence of a heterologous gene expressed in Wolbachia may be “codon optimized” to optimize gene expression based on the preferred codon usage in Wolbachia.
Nucleic acid is “operably linked” when it is placed into a functional relationship with another nucleic acid sequence. For example, DNA for a presequence or secretory leader is operably linked to DNA for a polypeptide if it is expressed as a preprotein that participates in the secretion of the polypeptide; a promoter or enhancer is operably linked to a coding sequence if it affects the transcription of the sequence; or a ribosome binding site is operably linked to a coding sequence if it is positioned so as to facilitate translation. Generally, “operably linked” means that the DNA sequences being linked are near each other, and, in the case of a secretory leader, contiguous and in reading phase. However, operably linked nucleic acids (e.g. enhancers and coding sequences) do not have to be contiguous. Linking is accomplished by ligation at convenient restriction sites. If such sites do not exist, the synthetic oligonucleotide adaptors or linkers are used in accordance with conventional practice. In embodiments, a promoter is operably linked with a coding sequence when it is capable of affecting (e.g. modulating relative to the absence of the promoter) the expression of a protein from that coding sequence (i.e., the coding sequence is under the transcriptional control of the promoter).
“Transformation” refers to the transfer of a nucleic acid molecule into a new carrier (e.g. Wolbachia cell or phage or prophage). In embodiments, the nucleic acid molecule may be a plasmid that replicates autonomously or it may integrate into the genome of the host organism. Host organisms containing the transformed nucleic acid molecule may be referred to as “transgenic” or “recombinant” or “transformed” organisms. A “genetically modified” organism (e.g. genetically modified arthropod) is an organism that includes a nucleic acid that has been modified by human intervention. Examples of a nucleic acid that has been modified by human intervention include, but are not limited to, insertions, deletions, mutations, expression nucleic acid constructs (e.g. over-expression or expression from a non-natural promoter or control sequence or an operably linked promoter and gene nucleic acid distinct from a naturally occurring promoter and gene nucleic acid in an organism), extra-chromosomal nucleic acids, and genomically contained modified nucleic acids.
“Transinfection” as used herein refers to extracting a microbe (either a pure extraction or mixed with other organisms or substances) from its natural host and then infecting an unnatural host with the extract. The recipient organism is then transinfected with a foreign microbe.
The term “male arthropod killing factor” or “male killing gene” refers to a gene or a factor encoded by the gene from bacteria which provide a function that is required and/or beneficial to produce the phenotype of male killing used by various, unrelated bacterial infections (e.g., Wolbachia).
The term “variant” or “derivative” as used herein refers to an amino acid sequence derived from the amino acid sequence of the parent protein having one or more amino acid substitutions, insertions, and/or deletions. For example, a “male arthropod killing factor variant” includes a male arthropod killing factor that may have a number of amino acid changes. In some embodiments, the variants may be greater than about 20%, greater than about 30%, greater than about 40%, greater than about 50%, greater than about 60%, greater than about 70%, greater than about 80%, greater than about 90%, or greater than about 95%, identical to the parent nucleic acid sequence or amino acid sequence.
The term “about” as used herein when referring to a measurable value such as an amount, a percentage, and the like, is meant to encompass variations of ±20%, ±10%, ±5%, or ±1% from the measurable value.
Male Arthropod Killing Factors and Methods of Use
In some aspects, disclosed herein is a genetically modified arthropod, said arthropod comprising:
-
- a gene encoding a male arthropod killing factor; and
- a promoter operably linked to the gene encoding the male arthropod killing factor;
- wherein the expression of the gene encoding the male arthropod killing factor in arthropod embryos causes a reduction in viable surviving male offspring in comparison to arthropod embryos not expressing the gene encoding the male arthropod killing factor.
In some embodiments, the gene encoding the male arthropod killing factor is from a bacterium. In some embodiments, the gene encoding the male arthropod killing factor is from a phage or prophage. In some embodiments, the gene encoding the male arthropod killing factor is from Wolbachia (or prophage WO or phage WO). In some embodiments, the male arthropod killing factor is wmk (WD0626) or its homologs. In some embodiments, the male arthropod killing factor comprises the amino acid sequence SEQ ID NO:1, or a variant thereof.
In some embodiments, the reduction in viable male offspring is greater than 10% (for example at least 10%, at least 20%, at least 30%, at least 40%, at least 50%, at least 60%, at least 70%, at least 80%, at least 90%, at least 100%).
In some embodiments, the arthropod is an insect. In some embodiments, the insect is selected from the genera consisting of Aedes, Culex and Anopheles. In some embodiments, the insect is selected from the group consisting of Aedes albopictus, Aedes aegypti and Aedes polynesiensis. In some embodiments, the insect is Drosophila suzukii.
In some aspects, disclosed herein is a method for controlling a population of target arthropods, comprising:
-
- providing a gene encoding a male arthropod killing factor, and a promoter operably linked to the gene encoding the male arthropod killing factor;
- transforming a population of arthropods with the gene encoding the male arthropod killing factor and the promoter operably linked to the gene encoding the male arthropod-killing factor; and
- releasing the population of arthropods amongst a population of target arthropods, wherein the release of the arthropods reduces the population of target arthropods.
In some aspects, disclosed herein is a method for controlling a population of target arthropods, comprising:
-
- providing a gene encoding a male arthropod killing factor, and a promoter operably linked to the gene encoding the male arthropod killing factor;
- genetically transforming a bacteria, phage, or prophage with the gene encoding the male arthropod killing factor operably linked to the promoter;
- transinfecting a population of arthropods with the bacteria, phage, or prophage; and
- releasing the population of arthropods amongst a population of target arthropods, wherein the release of the arthropods reduces the population of target arthropods.
In some embodiments, the gene encoding the male arthropod killing factor is from a bacterium. In some embodiments, the gene encoding the male arthropod killing factor is from a phage or prophage. In some embodiments, the gene encoding the male arthropod killing factor is from Wolbachia (or prophage WO or phage WO). In some embodiments, the male arthropod killing factor is wmk (WD0626) or its homologs. In some embodiments, the male arthropod killing factor comprises the amino acid sequence SEQ ID NO:1, or a variant thereof. In some embodiments, the reduction in viable male offspring is greater than 10% (for example at least 10%, at least 20%, at least 30%, at least 40%, at least 50%, at least 60%, at least 70%, at least 80%, at least 90%, at least 100%, or more).
In some embodiments, the arthropod is an insect. In some embodiments, the insect is selected from the genera consisting of Aedes, Culex and Anopheles. In some embodiments, the insect is selected from the group consisting of Aedes albopictus, Aedes aegypti and Aedes polynesiensis. In some embodiments, the insect is Drosophila suzukii.
In some embodiments, the method further comprises providing an additional method of arthropod control. In some embodiments, the additional method of arthropod control is a sterile insect technique (SIT). In some embodiments, the additional method of arthropod control is an incompatible insect technique (IIT).
In some embodiments, disclosed herein is a genetically modified arthropod, said arthropod comprising: a gene encoding a male arthropod killing factor (or with cofactors) or a variant thereof. In some embodiments, the gene encoding the male arthropod killing factor is a heterologous gene.
In some embodiments, the male arthropod killing factor is wmk (WD0626). In some embodiments, the male arthropod killing factor is a homolog of wmk (WD0626). In some embodiments, the male arthropod killing factor is selected from the sequences disclosed in Table 1. In some embodiments, the gene encoding the male arthropod killing factor is selected from the sequences disclosed in Table 2A and 2B. For example, the male arthropod killing factor can be selected from the group consisting of wRec 0560, wInn homolog of WD0626, wBor homolog of WD0626, wMel WD0255, wMel WD0623, wInn homolog of WD0623, wBor homolog of WD0623, wMel WD0508, wMel WD0622, wInn homolog of WD0622, wBor homolog of WD0622, and wBif homolog of WD0622. In some embodiments, the male arthropod killing factor can be selected from the accession numbers WP_010962718.1, WP_038198911.1, WP_010962465.1, WP_010962717.1, WP_010962645.1, or WP_010962716.1.
In some embodiments, the wmk homolog fulfills at least one of the following: 1) sequence similarity of 40% or more to wmk; 2) genes with one or more HTH domain(s) in phage WO gene regions; and/or 3) HTH-containing genes present in the same phage gene region as any cif/cid/cin (genes WD0631 and WD0632 in strain wMel) homologs.
In some embodiments, the male arthropod killing factor comprises the amino acid sequence SEQ ID NO:1. In some embodiments, the male arthropod killing factor comprises the amino acid sequence SEQ ID NO:1, or a variant thereof.
In some embodiments, the gene encoding the male arthropod killing factor comprises the amino acid sequence encoded by the nucleic acid sequence SEQ ID NO:2. In some embodiments, the gene encoding the male arthropod killing factor comprises the amino acid sequence encoded by the nucleic acid sequence SEQ ID NO:2, or a variant thereof.
In some embodiments, the gene encoding the male arthropod killing factor sequence has been codon optimized.
In some embodiments, the male arthropod killing factor comprises an amino acid sequence that is at least 30% identical (for example, at least 30%, at least 40%, at least 50%, at least 60%, at least 70%, at least 80%, at least 90%, at least 95%, at least 99%) to the amino acid sequence SEQ ID NO:1.
In some embodiments, the gene encoding the male arthropod killing factor comprises a sequence that is at least 30% identical (for example, at least 30%, at least 40%, at least 50%, at least 60%, at least 70%, at least 80%, at least 90%, at least 95%, at least 99%) to the amino acid sequence encoded by the nucleic acid sequence SEQ ID NO:2.
In some embodiments, the male arthropod killing factor comprises an amino acid sequence that is at least 20% identical to the amino acid sequence SEQ ID NO:1. In some embodiments, the gene encoding the male arthropod killing factor comprises a sequence that is at least 20% identical to the amino acid sequence encoded by the nucleic acid sequence SEQ ID NO:2.
In some embodiments, the male arthropod killing factor comprises the amino acid sequence SEQ ID NO:1, or a fragment thereof. In some embodiments, the male arthropod killing factor comprises an amino acid sequence encoded by the nucleic acid sequence SEQ ID NO:2, or a fragment thereof.
In some embodiments, the male arthropod killing factor comprises an amino acid sequence at least 60% (for example, at least 60%, at least 65%, at least 70%, at least 75%, at least 80%, at least 85%, at least 90%, at least 95%, at least 96%, at least 97%, at least 98%, at least 99%) identical to SEQ ID NO: 1.
In some embodiments, the male arthropod killing factor comprises an amino acid sequence encoded by a nucleic acid sequence at least 60% (for example, at least 60%, at least 65%, at least 70%, at least 75%, at least 80%, at least 85%, at least 90%, at least 95%, at least 96%, at least 97%, at least 98%, at least 99%) identical to SEQ ID NO: 2.
In some embodiments, the male arthropod killing factor comprises a wmk homolog listed in Table 1, or a variant or fragment thereof. In some embodiments, the male arthropod killing factor comprises a sequence at least 60% (for example, at least 60%, at least 65%, at least 70%, at least 75%, at least 80%, at least 85%, at least 90%, at least 95%, at least 96%, at least 97%, at least 98%, at least 99%) identical to a wmk homolog listed in Table 1, or a variant or fragment thereof.
In some embodiments, the male arthropod killing factor comprises a wmk homolog listed in Table 2A, or a variant or fragment thereof. In some embodiments, the male arthropod killing factor comprises a sequence at least 60% (for example, at least 60%, at least 65%, at least 70%, at least 75%, at least 80%, at least 85%, at least 90%, at least 95%, at least 96%, at least 97%, at least 98%, at least 99%) identical to a wmk homolog listed in Table 2A, or a variant or fragment thereof.
In some embodiments, the male arthropod killing factor comprises a wmk homolog listed in Table 2B, or a variant or fragment thereof. In some embodiments, the male arthropod killing factor comprises a sequence at least 60% (for example, at least 60%, at least 65%, at least 70%, at least 75%, at least 80%, at least 85%, at least 90%, at least 95%, at least 96%, at least 97%, at least 98%, at least 99%) identical to a wmk homolog listed in Table 2B, or a variant or fragment thereof.
In some embodiments, the male arthropod killing factor comprises a wmk homolog listed in Table 3, or a variant or fragment thereof. In some embodiments, the male arthropod killing factor comprises a sequence at least 60% (for example, at least 60%, at least 65%, at least 70%, at least 75%, at least 80%, at least 85%, at least 90%, at least 95%, at least 96%, at least 97%, at least 98%, at least 99%) identical to a wmk homolog listed in Table 3, or a variant or fragment thereof.
In some embodiments, the male arthropod killing factor comprises a wmk homolog listed in Table 6, or a variant or fragment thereof. In some embodiments, the male arthropod killing factor comprises a sequence at least 60% (for example, at least 60%, at least 65%, at least 70%, at least 75%, at least 80%, at least 85%, at least 90%, at least 95%, at least 96%, at least 97%, at least 98%, at least 99%) identical to a wmk homolog listed in Table 6, or a variant or fragment thereof.
In some aspects, disclosed herein is a genetically modified arthropod, said arthropod comprising: a gene encoding a male arthropod killing factor (or a variant thereof); and a promoter operably linked to the gene encoding a male arthropod killing factor or a variant thereof. In some embodiments, the promoter is heterologous. In some embodiments, the promoter is an arthropod promoter. In some embodiments, the gene encoding the male arthropod killing factor is inserted at an endogenous promoter in the arthropod. In some embodiments, the gene encoding the male arthropod killing factor is inserted at a heterologous promoter in the arthropod.
In some aspects, disclosed herein is a genetically modified bacterium comprising:
-
- a gene encoding a male arthropod killing factor or a variant thereof; and
- a promoter operably linked to the gene encoding a male arthropod killing factor;
- wherein the gene occurs at a non-naturally occurring genomic location in the bacterium.
In some aspects, disclosed herein is an arthropod infected with a bacterium, wherein the bacterium comprises:
-
- a gene encoding a male arthropod killing factor; and
- a promoter operably linked to the gene encoding the male arthropod killing factor;
- wherein the gene encoding the male arthropod killing factor occurs at a non-naturally occurring genomic location in the bacterium.
In other aspects, disclosed herein is a method for controlling a population of target arthropods, comprising:
-
- providing a genetically modified bacterium comprising:
- a gene encoding a male arthropod killing factor, and a promoter operably linked to the gene encoding the male arthropod killing factor;
- infecting a population of replacement arthropods with the genetically modified bacterium; and
- releasing the replacement arthropods amongst a population of target arthropods, wherein the release of the replacement arthropods reduces the population of target arthropods.
In some embodiments, the gene encoding the male arthropod killing factor in arthropod embryos causes a reduction in viable surviving adult male offspring in comparison to arthropod embryos not expressing the gene encoding the male arthropod killing factor.
In addition to genetically bacteria, the methods herein can also use genetically modified phage or prophage. The genetically modified bacteria, phage, or prophage can be transinfected into a new host (a non-natural host). The genetically modified hosts can use any number of promoters known in the art to drive gene expression. The genetically modified hosts can comprise the WD0626 gene, or homologs thereof, or can also include additional male killing genes or cofactors in combination with the WD0626 gene. The genetically modified hosts can comprise multiple copies of the WD0626 gene, or multiple copies of homologs thereof, or any combinations thereof. In addition, as disclosed herein, the various methods of controlling arthropods can be used individually or can be used in various combinations to employ multiple methods in succession, or simultaneously, in methods of controlling a population of target arthropods.
Further disclosed herein are genetically modified bacteria, genetically modified phage, and/or genetically modified prophage for controlling and/or reducing populations of arthropods. Also disclosed herein is an arthropod transinfected with a male-killing microbe, phage or bacteria that naturally infects another host.
Methods in Combination with the Sterile Insect Technique (SIT) and/or an Incompatible Insect Technique (IIT)
In some embodiments, the male arthropod killing factors and methods of use disclosed herein can be used in combination with an additional arthropod (for example, mosquito) control technique. In some embodiments, the additional arthropod control technique is selected from a sterile insect technique (SIT) or an incompatible insect technique (IIT).
In some embodiments, the male arthropod killing factors and methods of use disclosed herein can be used in combination with a Sterile Insect Technique (SIT). The concept of the sterile insect technique (SIT) was first discovered by Knipling in 1955 (Knipling, E. F. J Econ Entomol 48, 459-462 (1955)). SIT is the use of sterile males to suppress populations of insects. SIT works by periodic controlled releases of vast numbers of sterile male insects into wild populations. In principle, these sterile males outnumber and outcompete wild males for matings with wild females. If a female mates with a sterile male she will lay eggs that do not hatch. If the proportion of sterile males consistently exceeds the proportion of fertile males then each new generation's reproduction is suppressed. As the wild population numbers dwindle, SIT becomes more and more effective creating a negative feedback loop that ultimately eradicates the species in an area. One major advantage of SIT population suppression versus traditional insecticide treatment is that it is species specific and environmentally safe. Three major processes are necessary for the implementation of SIT: 1) a method of sterilization; 2) a method of sex separation; and 3) a method of dispersal.
The historical example of SIT is Knipling's and the USDA's rearing of irradiated sterile males to eradicate the New World Screwworm (Cochliomya hominivorax) in North America and Mexico (Bushland, R. C., et al. Science 122, 287-288 (1955)). Screwworm is a deadly livestock pest which causes myiasis (an infestation of parasitic fly larvae that feed on host tissues) (Lindquist, D. A., et al. Med Vet Entomol 6, 2-8 (1992)). Initial field tests were carried out in Florida starting in 1951 and later in 1954 on the island of Curacao (Baumhover, A. H. et al. J Econ Entomol 48, 462-466 (1955)). This initial program utilized gamma rays of cobalt to sterilize male pupae (Bushland, R. C. & Hopkins, D. E. J Econ Entomol 44, 725-731 (1951)). Adult flies were then dispersed over the island by weekly release from an airplane. After 6 months of releases, screwworm was completely eradicated from the island (Baumhove. Ah. J Amer Med Assoc 196, 240 (1966)). Using the same technique, screwworm was eradicated from Florida and the Southeast USA by 1959 (Baumhove. Ah. J Amer Med Assoc 196, 240 (1966); Baumhover, A. H., et al. J Econ Entomol 52, 1202-1206 (1959)) and entirely from North and Central America by 1995 (Baumhover, A. H. Baumhover: A Personal Account of Screwworm Eradication. Pioneer Lecture presentation (1997)). SIT based eradication of the screwworm was later replicated in Libya (1990) when a shipment of contaminated livestock caused an outbreak; the technique has been proven to be a useful suppression tactic for many insects (Lindquist, D. A., et al. Med Vet Entomol 6, 2-8 (1992)).
The physical quality or “fitness” of sterile insects produced for SIT is of paramount importance for the application. One downside of canonical sterilization by irradiation is that many insects are not as resilient to this treatment as screwworm. For example, mosquitoes are more sensitive to irradiation and cannot be irradiated without significant fitness reductions and lethality (Benedict, M. Q. & Robinson, A. S. Trends Parasitol 19, 349-355 (2003); Dame, D. A., et al. Historical applications of induced sterilization in field populations of mosquitoes. Malaria J 8 (2009)). Thus, alternative means of sterilization are useful inventions for the development and application of SIT. These additional methods of inducing sterility in insects include chromosomal disruptions, chemical sterilization, and sex ratio distortion (Benedict, M. Q. & Robinson, A. S. Trends Parasitol 19, 349-355 (2003)).
The sterile insect technique involves methods to make male mosquitoes sterile using techniques such as irradiation, chemical sterilization, and/or genetic sterilization techniques. Radiation-sterilization, as used for example in conventional sterile insect technique, is an example of conditional (in this case inducible by irradiation) paternal-effect lethality; sterilization with chemosterilants, e.g. thiotepa, is another. Each of these approaches work by damaging the DNA in the sperm, thus degrading the genetic information that it carries to the point that many zygotes die. US20170188559 describes conditional lethal expression systems for insects, their use, and methods of population control using transformed insects. US20170188559 uses improved female specific, repressible, dominant, lethal genetic system, with earlier onset of the lethal effect. Thus, this method achieves a similar result to techniques such as the Sterile Insect Technique (SIT) in insects, without the problems associated with SIT, such as the cost, danger to the user, and reduced competitiveness of the irradiated organism.
In some embodiments, the male arthropod killing factors and methods of use disclosed herein can be used in combination with an incompatible insect technique (IIT). Cytoplasmic incompatibility (CI) is a conditional sterility induced by a secreted bacterial sperm toxin produced from Wolbachia infections in insect gonads. Hannes Laven was the first to pioneer research on Wolbachia as a tool for SIT. He described how Culex pipiens mosquito isolates were sterile when mated with isolates from different regions of Europe (Laven, H. Chapter 7: Speciation and Evolution in Culex pipiens. 251 (Elselvier, 1967)). Realizing the potential, Laven isolated a strain of Culex pipiens fatigans (major vector of filariasis) which would be sterile when mated to the same species in Burma. Unbeknownst to Laven, his mosquito strain was infected with a corresponding strain of Wolbachia incompatible with the wild type populations in Burma. Despite not understanding the functionality of the sterility, Laven was able to use Wolbachia sterilized male mosquitoes to eradicate populations of the local mosquito vector in Burma (Laven, H. Nature 216, 383 (1967)).
The IIT methods can use different methods of cytoplasmic incompatibility (CI)-based population suppression. Wolbachia bacteria can cause a form of conditional sterility, which can provide an alternative to genetic modification or irradiation. See Mains, J, et al. Female Adult Aedes albopictus Suppression by Wolbachia-Infected Male Mosquitoes. Sci Rep. 2016 Sep. 23; 6:33846. doi: 10.1038/srep33846. U.S. Pat. No. 7,868,222 is directed to methods for artificially infecting an Aedes mosquito which can be introduced into a mosquito population to control the reproduction capability of the population by introducing an incompatible Wolbachia infection. US2013/0259846 is directed to a formulation that may include artificially generated adult insect carriers of a larvicide in which the larvicide has minimal impact on the adult insect and which larvicide affects juvenile survival or interferes with metamorphosis of juvenile insects to adulthood.
WO/2017/214476, discloses methods of utilizing bacterial genes that induce cytoplasmic incompatibility (CI), and discloses the minimal molecular components from the Wolbachia genome that are sufficient to induce sterility by a transgenic means, independent of the Wolbachia bacterium. U.S. Pat. No. 9,090,911 describes a line of mosquito adapted by infection of variants of the Wolbachia strain wMel. See also, Lees, R. et al. Back to the future: the sterile insect technique against mosquito disease vectors. Current Opinion in Insect Science 2015, 10:156-162, which provides additional description of the SIT and IIT technologies.
After sterilization, male insects can then be separated from female insects, delivered to the target site, and released for mating with wild females to eradicate a pest population.
U.S. Pat. No. 9,125,388 describes whereby biological control of an insect is achieved by the release of a dominant negative lethal gene under the control of transcriptional regulators.
The publications describing the SIT and IIT methods above are hereby incorporated by reference in their entirety.
Methods in Combination with Population Replacement Strategies
In some embodiments, the male arthropod killing factors and methods of use disclosed herein can be used in combination with a Population Replacement Strategy (PRS). The goal of PRS is to replace wild pest or vector populations with those that are not competent to function as pests or vectors of human disease (Sinkins, 2004, Insect Biochem Mol Biol, 34, 723-9; Dobson, Brelsfoard and Dobson, 2009, AsPac J. Mol. Biol. Biotechnol., 17, 55-63). Population Replacement is dependent on two pieces of technology:
-
- 1) A beneficial trait that is desired in the target arthropod
- 2) A genetic drive mechanism to spread the desired trait through the arthropod population (Sinkins and Gould, 2006, Nat Rev Genet, 7, 427-35).
For example, WO/2017/214476, discloses methods of utilizing bacterial genes that induce cytoplasmic incompatibility (CI), which is a natural genetic drive mechanism used by various, unrelated bacterial infections (e.g., Wolbachia and Cardinium endosymbionts).
Other approaches involving population replacement in the control of disease vectors is the Eliminate Dengue project. This method uses the naturally occurring Wolbachia strain wMel to introduce both a desirable trait, the inhibition of mosquito vector competence for Dengue virus and other human pathogens (Walker et al., 2011, Nature, 476, 450-3; Aliota et al., 2016, PLoS Negl Trop Dis, 10, e0004677; Dutra et al., 2016, Cell Host Microbe), and the genetic drive mechanism of CI. This technique has had limited success in field trials, requires massive mosquito releases (Hoffmann et al., 2011, Hoffmann et al., 2014), and the horizontal transfer of Wolbachia into hosts that are frequently inhospitable to stable infection (Hughes et al., 2011, PLoS Pathog, 7, e1002043; Hughes et al., 2014, Proc Natl Acad Sci USA, 111, 12498-503).
Arthropods and Infectious Disease Vectors
The inventors have identified a primary mechanism for the male killing phenotype and disclose herein new methods for control of arthropod (for example, insects) pests and disease vectors, such as mosquitoes transmitting the Dengue fever and Zika viruses.
In one embodiment, the arthropod is an insect. In one embodiment, the arthropod is a mosquito. In one embodiment, the mosquito is selected from the genera consisting of Aedes, Culex and Anopheles. In one embodiment, the mosquito is an Aedes mosquito. In one embodiment, the mosquito is an Anopheles mosquito. In one embodiment, the mosquito is a Culex mosquito. In one embodiment, the Aedes mosquito species is selected from the group consisting of Aedes albopictus, Aedes aegypti and Aedes polynesiensis. In one embodiment, the Anopheles mosquito species is Anopheles gambiae. In one embodiment, the Culex mosquito species is Culex pipiens.
In one embodiment, disclosed herein are methods for controlling or reducing populations of insects that transmit human or veterinary pathogens. In one embodiment, the pathogen is selected from dengue virus, Zika virus, a malaria parasite (Plasmodium genus), West Nile virus, yellow fever virus, chikungunya virus, Japanese encephalitis, St. Louis encephalitis and Western and Eastern Equine Encephalitis viruses.
In one embodiment, disclosed herein are methods for controlling or reducing populations of insects that transmit trypanosomes including African sleeping sickness, Chagas disease, and Nagana. In one embodiment, the pathogen is Trypanosoma cruzi. In one embodiment, the pathogen is Trypanosoma brucei. In one embodiment, the insect is of the genus Glossina. In one embodiment, the insect is Glossina morsitans. In one embodiment, the insect is a Tsetse fly. In one embodiment, the insect is a kissing bug. In one embodiment, the insect is of the genus Rodnius. In one embodiment, the insect is Rhodnius prolixus.
In one embodiment, disclosed herein are methods for controlling or reducing populations of arthropods that transmit rickettsioses and pathogens within Anaplasmatacea including Rickettsias rickettsii, africae, parkeri, sibirica, conorii, slovaca, peacockii, philipii, rickettsii Hlp2, heilongjiangensis, japonica, montanensis, massiliae, rhipicephali, amblyommii, helvetica, monacensis, buchneri, hoogstralli, felis, akari, australis, canadensis, prowazekii, typhi, bellii. In one embodiment, the arthropod is a tick. In one embodiment, the arthropod is a tick of the genera Amblyomma, Ixodes, or Rhipicephalus. In one embodiment, the disease is epidemic typhus. In one embodiment, the disease is scrub typhus. In one embodiment, the disease is an Ehrlichiosis. In one embodiment, the pathogen is of the genus Ehrlichia. In one embodiment, the pathogen is of the genus Anaplasma. In one embodiment, the pathogen is of the genus Orientia. In one embodiment, the arthropod is a chigger of the genus Leptotrombidium. In one embodiment, the arthropod is a louse of the genus Pediculus. In one embodiment, the arthropod is a flea of the genus Pulex.
In one embodiment, disclosed herein are methods for controlling sandflies that transmit leishmaniasis. In one embodiment, the insect is of the genus Phlebotomus. In one embodiment, the pathogen is of the genus Leishmania. In one embodiment, the pathogen is Leishmania donovani, Leishmania infantum, or Leishmania Chagasi.
In one embodiment, the insect is of various aphids including: Acyrthosiphon kondoi, Brevicoryne brassicae, Rhopalosiphum maidis, Aphis gossypii, Aphis craccivora, Myzus persicae, Rhopalosiphum padi, Acyrthosiphon pisum, Rhopalosiphum rufiabdominalis, Metopolophium dirhodum, Aphis glycine, Therioaphis trifolii, Lipaphis erysimi, Rhopalosiphum padi.
In one embodiment, disclosed herein are methods for controlling the armyworm agricultural pest species including Leucania convecta, Spodoptera exempta, Spodoptera Mauritia, Spodoptera exigua, Mythimna separate, Leucania stenographa.
In one embodiment, disclosed herein are methods for controlling pests of beans and beets. In one embodiment, the insect is either the Bean fly (Ophiomyia phaseoli), the Bean leafroller (Omiodes diemenalis), the Bean looper or Mocis (Mocis alterna), the Bean podborer (Maruca vitrata), the Bean spider mite (Tetranychus ludeni), the Beet webworm (Spoladea recurvalis), the Large Brown bean bug (Riptortus serripes), the Small Brown bean bug (Melanacanthus scutellaris)
In one embodiment, disclosed herein are methods for controlling the Blue oat mite (Penthaleus major). In one embodiment, the invention is useful for controlling the Brown flea beetle (Chaetocnema sp.). In one embodiment, the invention is useful for controlling the Brown mind (Creontiades pacificus). In one embodiment, the invention is useful for controlling the Brown shield bug (Dictyotus caenosus). In one embodiment, the invention is useful for controlling the Brown wheat mite (Petrobia latens). In one embodiment, the invention is useful for controlling the Bruchid, Cowpea (Callosobruchus maculatus).
In one embodiment, disclosed herein are methods for controlling pests of Corn including: the Corn aphid (Rhopalosiphum maidis), and the Corn earworm (Helicoverpa armigera).
In one embodiment, the invention is useful for controlling pests of cotton including the Cotton aphid (Aphis gossypii), Cotton bollworm (Helicoverpa armigera), the Cotton harlequin bug (Tectocoris diophthalmus), the Cotton leafhopper (Amrasca terraereginae), the Cotton leafperforator (Bucculatrix gossypii), the Cotton looper (Anomis flava), the Cottonseed bug (Oxycarenus luctuosus), the Cotton seedling thrip (Thrips tabaci), the Cotton tipworm (Crocidosema plebejana), and the Cotton webspinner (Achyra affinitalis).
In one embodiment, disclosed herein are methods for controlling the Diamondback moth (Plutella xylostella). In one embodiment, the invention is useful for controlling the Dried fruit beetle (Carpophilus spp.). In one embodiment, the invention is useful for controlling the Eastern false wireworm (Pterohelaeus spp.). In one embodiment, the invention is useful for controlling the Etiella moth (Etiella behrii). In one embodiment, the invention is useful for controlling the False wireworm (Pterohelaeus and Gonocephalum spp.). In one embodiment, the invention is useful for controlling the Flea beetles, Brown and Redheaded (Chaetocnema and Nisostra sp.). In one embodiment, the invention is useful for controlling the Flower beetle (Carpophilus spp.).
In one embodiment, disclosed herein are methods for controlling various Grasshoppers and locusts including the Grasshopper, Wingless (Phaulacridium vittatum), the Locust, Australian plague (Chortoicetes terminifera), the Locust, Migratory (Locusta migratoria), the Locust, Yellow-winged (Gastrimargus musicus), the Locust, Spur-throated (Austracris (Noamdacris) guttulosa).
In one embodiment, the invention is useful for controlling the Greenhouse whitefly (Trialeurodes vaporariorum). In one embodiment, the invention is useful for controlling the Green peach aphid (Myzus persicae). In one embodiment, the invention is useful for controlling the Green mind (Creontiades dilutus). In one embodiment, the invention is useful for controlling the Green vegetable bug (Nezara viridula). In one embodiment, the invention is useful for controlling the Green stink bug (Plautia affinis). In one embodiment, the invention is useful for controlling the Grey cluster bug (Nysius clevelandensis). In one embodiment, the invention is useful for controlling the Helicoverpa species (armigera and punctigera).
In one embodiment, disclosed herein are methods for controlling planthoppers. In one embodiment, the insect is the small brown planthopper (Laodelphax striatellus). In one embodiment, the invention is useful for preventing the transmission of crop diseases like Rice White Stripe Virus. In one embodiment, the invention is useful for controlling vectors of plant pathogens.
In one embodiment, disclosed herein are methods for controlling the Jassids and various leafhoppers including the Leafhopper, cotton (Amrasca terraereginae), the Leafhopper, lucerne (Austroasca alfalfae), the Leafhopper, maize (Cicadulina bimaculata), the Leafhopper, vegetable (Austroasca viridigrisea).
In one embodiment, disclosed herein are methods for controlling the Loopers including the Looper, Brown pasture (Ciampa arietaria), the Looper, Castor oil (Achaea janata), the Looper, Cotton (Anomis flava), the Looper, Sugarcane (Mocis frugalis), the Looper, Soybean (Thysanoplusia orichalcea), the Looper, Tobacco (Chrysodeixis argentifera), the Looper, Vegetable (Chrysodeixis eriosoma).
In one embodiment, disclosed herein are methods for controlling various Thrip pests including the Onion Thrip (Thrips tabaci), the Cotton seedling Thrip (Thrips tabaci), the Maize Thrip (Frankliniella williamsi), the Plague Thrip (Thrips imaginis), the tobacco Thrip (Thrips tabaci), the Tomato Thrip (Frankliniella schultzei), the Western flower Thrip (Frankliniella orientalis)
In one embodiment, disclosed herein are methods for controlling various Mite pests including the Mite, Bean spider (Tetranychus ludeni), Mite, Brown wheat (Petrobia latens), Mite, Blue oat (Penthaleus major), Mite, Peanut (Paraplonobia spp.), Mite, Redlegged earth (Halotydeus destructor), Mite, Strawberry spider (Tetranychus lambi), and the Two-spotted mite (Tetranychus urticae).
In one embodiment, disclosed herein are methods for controlling various whitefly pests including the Greenhouse whitefly (Trialeurodes vaporariorum), the Silverleaf whitefly (Bemisia tabaci biotype B and Australian native AN), and the Silverleaf whitefly (Bemisia tabaci biotype Q).
In one embodiment, disclosed herein are methods for controlling various fruit pests. In one embodiment, the arthropod is from the genera Drosophila. In one embodiment, the arthropod is Drosophila suzukii. In one embodiment, the arthropod is Drosophila recens, Drosophila subquinaria, Drosophila innubila, or related Drosophila species. Drosophila suzukii, commonly called the spotted-wing drosophila, is a vinegar fly closely related to Drosophila melanogaster. Unlike its vinegar fly relatives who are primarily attracted to rotting or fermented fruit, D. suzukii attacks fresh, ripe fruit by laying eggs under the soft skin. The larvae hatch and grow in the fruit, destroying the fruit's commercial value. The pest particularly (but not limited to) infests cherries, apples, apricots, persimmons, tomatoes, blueberries, grapes, nectarines, pears, plums, peaches, figs, raspberries and strawberries. Although D. suzukii is native to Southeast Asia, the fruit pest has recently invaded North and Central America as well as Europe, where it is expanding rapidly. Effective management of this pest is a challenge owing to the wide host range and short generation time. Therefore, monitoring and controlling D. suzukii is of great economic importance. However, traps and baits containing for instance apple cider vinegar, which are typically used for attracting vinegar flies such as D. melanogaster, are less efficient for attracting and trapping D. suzukii. In one embodiment, the insect is the Mexican Fruit Fly (Anastrepha ludens). In one embodiment, the insect is the Mediterranean Fruit Fly (Ceratitis capitata). In one embodiment, the insect is of the genus Anastrepha, Bactrocera, or Ceratitis. In one embodiment, the insect is a tephritid.
In one embodiment, disclosed herein are methods for controlling various other agricultural pests including: the red-houldered leaf beetle (Monolepta australis), Native budworm (Helicoverpa punctigera), Native whitefly (Bemisia tabaci), Northern armyworm (Mythimna separata), Oat aphid (Rhopalosiphum padi), Onion thrip (Thrips tabaci), Pale cotton stainer bug (Dysdercus sidae), Pea aphid (Acyrthosiphon pisum), Pea blue butterfly (Lampides boeticus), Peanut mite (Paraplonobia spp.), Peanut scarab (Heteronyx spp.), Pea weevil (Bruchus pisorum), Pinkspotted bollworm (Pectinophora scutigera), Plague thrip (Thrips imaginis), Podsucking bugs (Nezara viridula), Redbanded shield bug (Piezodorus oceanicus), Redheaded flea beetle (Nisotra sp.), Redlegged earth mite (Halotydeus destructor), Redshouldered leaf beetle (Monolepta australis), Rice root aphid (Rhopalosiphum rufiabdominalis), Rose grain aphid (Metopolophium dirhodum), Rough bollworm (Earias huegeliana), Rutherglen bug (Nysius vinitor), Seed harvesting ants (Pheidole spp.), Scarab, Black sunflower (Pseudoheteronyx sp.), Scarab, Peanut (JPG, 20.4 KB) (Heteronyx sp.), Shoot flies (Atherigona sp.), Silverleaf whitefly (Bemisia tabaci biotype B and Australian native AN), Silverleaf whitefly (Bemisia tabaci biotype Q), Sitona weevil (Sitona discoideus), Solenopsis mealybug (Phenacoccus solenopsis), Sorghum midge (Stenodiplosis sorghicola), Sorghum head caterpillar (Cryptoblabes adoceta), Soybean leafminer (Porphyrosela aglaozona), Soybean looper (Thysanoplusia orichalcea), Soybean moth (Aproaerema simplexella), Spotted alfalfa aphid (Therioaphis trifolii), Spur-throated locust (Austracris (Nomadacris) guttulosa), Strawberry spider mite (Tetranychus lambi), Swarming leaf beetle (Rhyparida spp.), Tortrix (Epiphyasa postvittana), True wireworm (Agrypnus spp.), Vegetable weevil (Listroderes difficilis), Weed web moth (Achyra affinitalis), Whitegrub (Heteronyx spp.), Wingless cockroaches (Calolampra spp.), Wireworm, False (Pterohelaeus and Gonocephalum spp.), Wireworm, True (Agrypnus spp.), Yellow peach moth (Conogethes punctiferalis). In one embodiment, the insect is Heteronychus arator. In one embodiment, the insect is of the genus Amnemus. In one embodiment, the insect is of the genus Pheidole. In one embodiment, the invention is useful for controlling the Black field cricket (Teleogryllus commodus, T. oceanicus, Lepidogryllus parvulus), the Black field earwig (Nala lividipes), the Black leaf beetle (Rhyparida nitida), the Black sunflower scarab (Pseudoheteronyx sp.). In one embodiment, the invention is useful for controlling the Cowpea bruchid (Callosobruchus maculatus). In one embodiment, the invention is useful for controlling the Cricket, Black field (Teleogryllus commodus, T. oceanicus, Lepidogryllus parvulus). In one embodiment, the invention is useful for controlling the Crop mind (Sidnia kinbergi). In one embodiment, the invention is useful for controlling the Cutworm (Agrotis spp.). In one embodiment, the invention is useful for controlling the Cabbage moth (Plutella xylostella). In one embodiment, the invention is useful for controlling the Castor oil looper (Achaea janata). In one embodiment, the invention is useful for controlling the Click beetle (Agrypnus spp.). In one embodiment, the invention is useful for controlling the Clover springtail (Sminthurus viridis). In one embodiment, the invention is useful for controlling the Cluster caterpillar (Spodoptera litura). In one embodiment, the invention is useful for controlling the Cockroach, Wingless (Calolampra spp.). In one embodiment, the invention is useful for controlling the Common grass blue butterfly (Zizina labradus). In one embodiment, the invention is useful for controlling the Legume webspinner (Omiodes diemenalis). In one embodiment, the invention is useful for controlling the Light brown apple moth (Epiphyas postvittana). In one embodiment, the invention is useful for controlling Mocis trifasciata. In one embodiment, the invention is useful for controlling Pantydia spp. In one embodiment, the invention is useful for controlling the Lucerne crownborer (Zygrita diva). In one embodiment, the invention is useful for controlling the Lucerne flea (Sminthurus viridis). In one embodiment, the invention is useful for controlling the Lucerne leafhopper (Austroasca alfalfae). In one embodiment, the invention is useful for controlling the Lucerne leafroller (Merophyas divulsana). In one embodiment, the invention is useful for controlling the Lucerne seed wasp (Bruchophagus roddi). In one embodiment, the invention is useful for controlling the Lucerne seed web moth (Etiella behrii).
In one embodiment, disclosed herein are methods for controlling forestry and wildlife pests such as the emerald ash borer. In one embodiment, the insect is of the genus Agrilus or specifically Agrilus planipennis. In one embodiment, the invention is useful for pests of trees and lumber.
In one embodiment, disclosed herein are methods for controlling various arthropods including Adalia bipunctata (two-spotted lady beetle), other ladybug species/genera (Harmonia, Adalia decempunctata, Cadra cautella (and other Cadra moths), Ephestia kuehniella (and other Ephestia moths), Cordylochernes scorpioides (pseudoscorpion), Tribolium (flour beetles), Hypolimnas butterflies, Acraea butterflies, or Ostrinia moths.
EXAMPLESThe following examples are set forth below to illustrate the compositions, methods, and results according to the disclosed subject matter. These examples are not intended to be inclusive of all aspects of the subject matter disclosed herein, but rather to illustrate representative methods and results. These examples are not intended to exclude equivalents and variations of the present invention which are apparent to one skilled in the art.
Example 1. wmk is a Wolbachia Prophage Gene that Kills Male Drosophila EmbryosWolbachia are maternally-transmitted bacteria that infect almost half of all arthropod species and many nematode species. In arthropods, these bacteria often selfishly manipulate host reproduction to enhance the fitness of infected females, thereby facilitating their own transmission and spread through the host population. Despite significant impacts of Wolbachia on animal reproduction1, evolution2-5, and vector control6,7, the bacterial genes underlying most of these reproductive manipulations remain elusive. One such phenotype is male killing, where the sons of infected females are selectively killed. In this example, it is demonstrated that a single gene in the eukaryotic association module of prophage WO8 kills male Drosophila embryos. The gene, hereafter denoted WO male killing (wmk), causes male lethality when transgenically expressed in uninfected Drosophila melanogaster. Expression of wmk results in a female-biased sex ratio, reduced hatching of male embryos, and male-specific cytological defects during early embryonic development that are typical of Wolbachia-induced male killing. The discovery of wmk commences microbial genetic studies of male killing, highlights the significance of genes carried by mobile genetic elements in shaping selfish symbiont phenotypes, and provides compositions and methods of male killing in suppression or modification of pest and vector populations6,7,9.
Background
Wolbachia (Order Rickettsiales) infect an estimated 40-52% of all arthropod species and 47% of filarial nematode species, making them the most widespread intracellular bacterial symbiont in animals. Concentrated in host testes and ovaries, Wolbachia primarily transmit cytoplasmically from mother to offspring. In arthropod reproductive tissues and fertilized embryos, Wolbachia deploy cunning manipulations to achieve a greater proportion of transmitting females in the host population. Collectively, these strategies are categorized as reproductive parasitism.
Male killing, or selective death of an infected female's sons, is one such form of reproductive parasitism. It enhances the fitness of Wolbachia-infected females in three potential ways: (i) reducing brother-sister competition for limited resources, (ii) reducing inbreeding, and/or (iii) providing nutrients in cases where infected sisters cannibalize embryos of their dead brothers. Male-killing Wolbachia are widespread in several major insect orders and in pseudoscorpions. In addition, male-killing Spiroplasma, Rickettsia, and Arsenophonus occur in diverse hosts including flies, ladybugs, and wasps.
Male killing can have several significant impacts on host evolution. For example, male death may lead to host extinction or reduce the effective population size of the host. As a consequence, theory specifies that fixation of deleterious alleles in host populations is more likely, and fixation of beneficial alleles is conversely less likely. Male killing can also impose strong selection on hosts to counter the sex ratios shifts and lethality. Evolutionary outcomes include mate preference between uninfected males and females, a shift towards more mate-attracting behaviors by females or male mate choice, and suppression of the phenotype.
As they manipulate arthropod reproduction to drive through host populations, Wolbachia are currently used in two vector control strategies: population suppression to reduce the population size of mosquitoes and population replacement to transform mosquito populations that transmit pathogens to ones that cannot transmit pathogens. This is through the application of another parasitism phenotype, cytoplasmic incompatibility (CI), where most offspring of infected males and uninfected females are killed. Notably, population genetic modeling demonstrates that male killing can be deployed in conjunction with population suppression techniques to speed up eradication or reduction of a target arthropod population and increase the likelihood of success. However, the genetic basis of Wolbachia male-killing has remained a mystery for more than sixty years and the causative gene of the Spiroplasma male-killing phenotype has only recently been reported, so vector and pest control applications have yet to be experimentally validated. This is largely due to the inability to culture these obligate intracellular bacteria.
Results
Genomic Analysis of Male-Killing Gene Candidates
In this example, the inventors sought to determine the Wolbachia genetic basis of the male-killing phenotype. Notably, Wolbachia can be multipotent because some strains induce at least two reproductive parasitism phenotypes (e.g., CI and male killing) depending on the host background or environmental conditions. For example, the wRec strain of D. recens causes CI in its native host but kills males when introgressed into the genetic background of its sister-species, D. subquinaria. Importantly, wMel and wRec share 99.7% nucleotide identity, which raises the hypothesis that the CI-inducing wMel genome may also harbor male killing genes. Although wMel is not known to naturally cause male killing, it is of interest because it is the natural strain of the only host that is genetically tractable and is so closely-related to a natural male killer, making it the best current system to test Wolbachia gene candidates for the phenotype. A long-standing question is whether multipotency is due to the same gene(s) expressing different reproductive parasitism phenotypes or alternatively if different genes underpin the various forms of reproductive parasitism. Several reproductive parasitism gene candidates were previously assessed for both male killing and CI, including cifA and cifB, and were ruled out their involvement in male killing.
There are several expectations for a putative Wolbachia male-killing gene. First, transgenic expression should recapitulate the embryonic cytological defects typically induced by male killing. Second, Wolbachia expression of the candidate gene should occur by the time male death naturally occurs in a given host. Third, a male-killing Wolbachia gene should be shared across male-killing strains but not necessarily absent from strains not known to cause male killing. In other words, the gene may be more common than the phenotype because hosts commonly develop resistance to male killing, presumably due to the strong evolutionary pressure to avoid extinction. Indeed, as previously mentioned, Wolbachia can induce either male killing or CI in different hosts or rearing conditions, which may be related to resistance in some hosts. Fourth, if there is a single gene that causes male killing in most or all cases, then the gene may rapidly evolve due to natural selection in diverse host backgrounds that suppress male killing. In this example, based on genomic analyses, transgenic expression, and cytological characterizations, a prophage WO gene from wMel Wolbachia of Drosophila melanogaster that causes male killing is identified.
To generate a shortlist of male-killing gene candidates, the following criteria were used: (i) universal presence in the genomes of male-killing strains wBif from D. bifasciata, wInn from D. innubila, wBor from D. borealis, wRec from D. recens as well as the CI strain nearly identical to wRec, wMel from D. melanogaster; (ii) genomic location in at least one prophage WO region because parasitic Wolbachia all have intact or remnant prophage WO regions with eukaryotic association module genes, and the two previous parasitism genes, cifA and cifB, are both prophage genes, making it likely that other parasitism genes share a phage origin; (iii) exclusion of highly repetitive elements, including insertion sequence elements, reverse transcriptases of group II intron origin, and large serine recombinases that likely facilitate phage WO lysogeny; and (iv) exclusion of disrupted genes (e.g., early stop codons) in one or more strains.
Table 1 shows seven genes from Wolbachia that fit these criteria. One of these genes, cifA, was previously evaluated by transgenic expression, and it did not exhibit a biased sex ratio. Others include a predicted ankyrin repeat (WD0550), two Rpn genes (recombination-promoting nucleases WD0297, WD0627), Phospholipase D (WD1243), and a hypothetical protein (WD0628). The remaining gene, WD0626, was a gene identified in multi-omic analysis for CI genes. This gene, hereafter denoted wmk for WO-mediated killing, is a putative transcriptional regulator in prophage WOMelB that is predicted to encode two helix-turn-helix (XRE family) DNA-binding domains (NCBI conserved domains E=5.9×10−11. E=6.5×10−10). wmk in wMel has a single amino acid difference relative to its homolog in wRec. Therefore, it was assessed for putative male killing.
The wmk Gene is Common and Found in all Sequenced Male-Killing Genomes
Phylogenetic analyses indicate that wmk homologs are common in phage WO-containing Wolbachia including the above-mentioned male-killing strains (
wmk (locus WD0626 in the wMel Wolbachia genome of Drosophila melanogaster) is a putative transcriptional regulator in prophage WOMelB that encodes two putative helix-turn-helix (HTH_XRE) DNA-binding domains (NCBI conserved domains E=5.9×10−11, E=6.5×10−10). Comparative genomic studies originally identified wmk and its homologs as candidates for inducing cytoplasmic incompatibility (CI)10-12, which is a common type of reproductive parasitism whereby crosses between infected males and uninfected females result in embryonic lethality. Rescue of this lethality occurs if the female and her eggs are infected with a compatible Wolbachia strain. CI, however, was later attributed to the nearby prophage WO genes cifA and cifB10,13. Given that CI-causing Wolbachia can be multipotent and also cause male killing, depending on the host and environmental conditions14-16, wmk was functionally interrogated to assess its potential involvement in male killing. Notably, wmk in wMel has a single amino acid difference compared to its homolog in the wRec strain17 that is capable of male killing when moved to a non-native hose15. Moreover, these two genomes share 99.7% nucleotide identity.
Expression of wmk in Fruit Flies Causes a Female-Biased Sex Ratio
Transgenic D. melanogaster flies were generated that express codon-optimized wmk (codons changed to those preferentially used by Drosophila) with the Gal4-UAS expression system. Three other transgenes were also evaluated: WD0625 in prophage WO that encodes a putative MPN/Mov34/PAD-1 metalloprotease domain (DUF2466, NCBI conserved domain E=3.85×10−41), WD0508 in the prophage WO-associated Octomom region that is another predicted transcription regulator with two HTH_XRE DNA-binding domains (NCBI conserved domains E=1.70×10−9, E=1.99×10−11, a homolog of wmk), and WD0034, a hypothetical non-phage gene that is hereafter labeled ‘control gene’ and shares a transgenic insertion site with wmk. These three genes were previously described and do not recapitulate CI10. In the experiments below, these transgenes were expressed within heterozygotes under the control of an Act5c-Gal4 driver, which leads to strong, ubiquitous transgene expression beginning with zygotic transcription ˜2 h after egg deposition (aed) and continues through adulthood. Genetic crossing schemes for all experiments are described in the methods.
To assess potential male killing, adult sex ratios were first quantified in gene-expressing (Act5c-Gal4; UAS-wmk) flies. wmk transgene expression beginning in early embryogenesis of uninfected D. melanogaster results in a significant reduction in the average male:female sex ratio (number of males/number of females) to 0.65, suggesting a 35% reduction in gene-expressing males (
The wmk-induced change in sex ratio is not consistent with other forms of reproductive parasitism. Results are not CI-like because (i) CI is not known to have a sex ratio bias10, (ii) the phenotype begins long after hallmark CI defects such as delayed histone deposition19 and (iii) an infected background does not rescue the wmk phenotype, as would be observed if the effect were related to CI (
Expression of wmk in Fruit Flies Recapitulates Embryonic Death and Cytological Defects
Wolbachia-induced male killing typically occurs during embryogenesis in Drosophila20 To assess if wmk-induced male lethality is embryonic, the proportion of all embryos that hatch into larvae were examined (note only half of embryos express the transgene, see methods). The mean hatch rate from the wmk transgenic line (79.5%) was significantly lower than the mean hatch rate for control WD0034 embryos (91%) and WT embryos (86.4%) (
Typical cytological defects associated with Wolbachia-induced male killing in D. bifasciata span abnormal nuclei distribution, chromatin bridging, and pyknosis in male embryos and they begin largely at the time of host embryonic cellularization, which occurs ˜2.5 h aed20. To determine if wmk transgene expression in D. melanogaster also causes these same defects, DNA was stained with propidium iodide in wild type (WT) embryos and embryos expressing either wmk or the control transgene. The defects in embryos were then monitored (only half the embryos are expected to express the transgene, see methods). Several different defects were observed among embryos (
To determine if the defects in embryos resulted in visible abnormalities later in development, sibling embryos were fixed in a late stage of embryonic development, 16-17 h aed. There was a significant increase in necrotic embryos (embryos with cloudy staining from degraded DNA and lack of distinct nuclei) in wmk-associated offspring compared to controls. One category of necrotic embryos had no visible cephalic furrow or segmentation similar to unfertilized embryos (
To demonstrate that the cytological defects are male-specific, fluorescent in situ hybridization (FISH) was performed on embryos with a DNA probe specific to the Y chromosome (
wmk is Expressed in Drosophila Embryos Infected with Wolbachia
To establish a native gene expression profile of wmk, relative transcript abundance was measured in Wolbachia-infected embryos fixed 4-5 h aed (estimated time of death). In wMel-infected embryos, native wmk and control gene transcripts were approximately 10-fold lower than the highly expressed CI gene, cifA (
wmk homologs are common in phage WO-containing Wolbachia including the male-killing strains wInn (D. innubila)21, wBor (D. borealis)22 wRec (D. recens, causes male killing in non-native host)15, wBol (Hypolimnas bolina, causes CI when male killing is suppressed)3,23, and wCauB (Cadra cautella, causes male killing in non-native host)16 (
The relationship between multipotent Wolbachia genotype and phenotype is complex, but some patterns emerge. First, although transgenic wmk induces a male-killing phenotype in D. melanogaster, its native Wolbachia strain, wMel, induces CI10. Notably, wRec causes male killing when transferred to its sister species, D. subquinaria15. This is also known to occur in the opposite direction, where suppression of a male killer can lead to CI induction, indicating that male killing and CI strains may be able to switch back and forth between the two phenotypes23. Given that wRec and wMel and their wmk homologs are so closely related, it is possible that wMel can also phenotype switch from CI to male killing depending on the host. Second, and more generally across Wolbachia strains, while CI genes and phenotype often correlate, wmk is not always associated with male killing. For example, while CI-capable strains wMel and wRec have cifA and cifB genes, several strains that are not known to cause CI, including wInn, have unusual cifB homologs with stop codons in the middle of the gene, suggesting they are unable to induce CI (
While around a third of gene-expressing males are killed, transgene expression of wmk in D. melanogaster does not kill all gene-expressing males even though complete male death is typical in the wild. This could be for several reasons. First, the single amino acid difference between the protein sequences of wMel wmk and wRec wmk may account for the discrepancy. Second, another gene may additively contribute to male killing. This possibility has not been ruled out, but the most likely candidate, WD0625, does not appear to be involved. A third possibility is that expression may need to be at a different time, however, neither an egg-loading driver nor the ubiquitously and early-expressed Act5c-Gal4 driver leads to total male death. Fourth, variation in expression levels among individuals may be a factor. Finally, several host species are resistant to male killing3,15. It is possible that D. melanogaster is not as susceptible to male killing as other hosts like D. subquinaria, and it is only through high levels of transgene expression that males die. This could be why high levels of wmk expression are required for male death in this system, but not necessarily in natural cases of male killing (
The Wmk Protein is a Putative DNA-Binding Protein
Phyre 2 protein modeling was used to predict that Wmk from wMel is globular and composed of α-helical secondary structures matching several transcriptional regulators, suppressors, and DNA-binding proteins (
In addition, the amino acid divergence across homologs of Wmk including the wBif homolog and all homologs are shown in
Discussion
This example reports five key results showing wmk is a phage WO male-killing gene: (i) The gene occurs on a shortlist of candidate phage WO genes in Wolbachia male-killers, including the highly reduced phage WO genome of wRec and the divergent phage WO genome of wBif. (ii) wmk is common, divergent in sequence, and located in the eukaryotic association module of phage WO that is enriched with genes of eukaryotic function and homology. In this region, wmk is a few genes away from the two causative cytoplasmic incompatibility genes, cifA and cif/3. (iii) Transgenic expression of wmk consistently induces a male-killing-like phenotype spanning male-biased, early embryonic death coupled with expression in embryos and canonical cytological defects. (iv) Wink's predicted structure is conserved across arthropod hosts despite sequence divergence. (v) Finally, no other tested transgenes result in this phenotype, and wmk does not fit the profile of any other known form of reproductive parasitism.
The analyses herein indicate that Wink is a putative DNA-binding transcriptional regulator (
There is considerable amino acid sequence divergence in Wink homologs across several arthropod orders that harbor male-killing Wolbachia. One potential reason for the divergence is that if it a single gene kills many or all of these hosts in nature, it may be divergent due to selection to target the varied genetic and cellular bases of sex determination in these hosts. Second, if there is a single gene behind the phenotype, it could explain the relatively high frequency of host resistance since hosts would only have to counter-adapt to one gene product rather than multiple products. Under antagonistic coevolution, wmk would evolve to kill males, the host would adapt to resist the male killing, and wmk would follow suit and adapt again, continuing the evolutionary arms race. Third, in addition to coevolutionary bouts of wmk adaptation and host counter-adaptation, pleiotropy or neutral evolution of wmk could also explain the sequence divergence in wmk homologs, especially in hosts that do not exhibit male killing. In addition, there is difference in copy number of homologs across genomes, however, wmk is the most likely candidate as wRec only contains one homolog and it is most similar to wmk rather than others in the wMel genome.
Identification of male-killing genes have relevance to translational applications in pest or vector control as male killing can be used in population suppression to crash target populations. The use of male killing in conjunction with other population-crashing techniques like Sterile Insect Technique (SIT), where sterilized males are released to compete with fertile males, could decrease the time to crash the population and increase the chances of success. In this context, male killing genes can be used to transform an endosymbiotic microbe or host to either add or enhance male-killing ability. Alternatively, a male-killing infection could be established in a host where one does not natively exist. These techniques may be desirable in cases of invasive species of disease-carrying mosquitoes or agricultural pests. Techniques like SIT can fail if males are not completely sterile or because of reduced mating competitiveness with fertile males. Therefore, a two-pronged approach to simultaneously reduce viable matings in the wild (SIT) while killing off males (male-killing) could be used to effectively crash populations prone to SIT failure on their own.
The discovery of wmk-induced male killing brings a new understanding to the types of prophage genes that can interact with animal reproduction. Male-specific lethality occurs in many arthropods and has important influences on arthropod evolution3,15,27-30, such as modifying mate choice and selecting for male resistance to the phenotype31,32. Male killing may also serve as a means to speed up other population suppression methods for vectors or pests9. Thus, deciphering male-killing genes helps inform the crosstalk between reproductive parasites and their animal hosts as well as their potential efficacy in arthropod control programs6,9.
Materials and Methods
Comparative Genomics and Evolutionary Analysis
Putative Wmk domains were identified by a CD-SEARCH of NCBI's Conserved Domain Database (https://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi). For the full-length wmk analysis (
Wolbachia Gene Sequencing:
The Wolbachia genome of D. innubila (wInn) was provided by R. Unckless. The Wolbachia genomes of D. bifasciata (wBif) and D. borealis (wBor) were provided by F. Jiggins. The whole genomes will be published by the respective contributors at a later date, and only the gene regions involved in this publication are currently publicly available (the regions in
The D. innubila Wolbachia genome was sequenced from a single wild-caught female. Briefly, D. innubila were captured at the Southwest Research Station over baits consisting of store-bought white button mushrooms (Agaricus bisporus). DNA was extracted using the Qiagen Gentra Puregene Tissue kit (#158689, Germantown, Maryland, USA). A genomic DNA library was constructed for several individuals using a modified version of the Nextera DNA Library Prep kit (#FC-121-1031, Illumina, Inc., San Diego, CA, USA) reagents33. DNA from an infected female was sequenced on a fraction of an Illumina HiSeq 2500 System Rapid-Run to generate 14873460 paired-end 150 base-pair reads. Reads were aligned to a draft D. innubila genome and all non-aligned reads were assembled de novo using Spades34. Those contigs blasting to other Wolbachia accessions were retained as putative Wolbachia genomic contigs.
The Wolbachia genomes of wBif and wBor were sequenced from D. bifasciata (line bif-F-MK35) and D. borealis (line PG05.1636) respectively. Following the protocol developed in Ellegaard et al.37, Wolbachia cells were purified from ˜20 freshly laid (less than 2 hours) and bleach-dechorionated embryos by homogenizing them in phosphate-buffered saline solution (PBS) and conducting a series of centrifugation/filtration steps as explained in Ellegaard et al37. A multiple-displacement amplification was carried out directly on the bacterial pellet using the Replig midi kit (Qiagen). The amplified DNA was cleaned with QIAamp DNA mini kit (Qiagen). From each sample, both 3 kb mate-pair and 50 bp paired-end DNA libraries were prepared and sequenced on a 454 Roche FLX (Department of Biochemistry, Cambridge, UK) and Illumina HiSeq2000 instruments (The Genome Analysis Center, Norwich, UK) respectively. The sequencing generated 203,565 and 239,485 454 mate-pair reads as well as 35,415,012 and 30,624,138 Illumina reads for wBif and wBor respectively. De novo hybrid assemblies combining 454 reads and a 10% subset of the Illumina reads were performed in Newbler (454 Life Sciences Corp., Roche, Branford, CT 06405, US). Contigs blasting to other Wolbachia accessions were retained as putative Wolbachia genomic contigs. Scaffolds were extended to fill regions with “N”s using GapFiller v.1-1138.
Drosophila Strains:
The Wolbachia transgene strains were generated as described previously10. WD0626 (wmk) and WD0034 (control gene) were both inserted into an attP site in the BSC8622 (WT) line of genotype y1w67c23; P[CaryP]P2 obtained from the Bloomington Drosophila Stock Center. WD0625 was inserted into the BSC9723 strain, with a genotype of y1M[vas-int.Dm]ZH-2A w*; PBac[y+-attP-3B]VK00002. WD0508 was inserted into the y1M[vas-int.Dm]ZH-2A w*; P[CaryP]attP40 line. The Act5c-Ga14/CyO driver line is the same background as BSC25374, which is y1w*; P[Act5C-GAL4-w]E1/CyO. The maternal triple driver (MTD) strain BSC31777, genotype P[w[+mC]=otu-GAL4::VP16.R]1, w[*];P[w[+mC]=GAL4-nos.NGT]40; P[w[+mC]=GAL4::VP16-nos.UTR]CG6325[MVD1], was provided by J. Nordman. The expression experiments were done using y1w* flies to measure native Wolbachia gene expression. The nanos-Gal4 strain used in
Drosophila Rearing:
D. melanogaster were reared on 4% cornmeal (w/v), 9% molasses (w/v), 1.6% yeast (w/v) (CMY) media. The flies developed at 25° C. at 80% humidity with a 12 h light/dark cycle. Virgin flies were stored at room temperature after collections. During virgin collections, stocks were maintained at 25° C. during the day and at 18° C. at night. Wolbachia-uninfected transgene or driver lines were generated via tetracycline treatment of infected lines as described previously10.
Crossing Scheme: Experimental Design
Most Drosophila experiments (unless otherwise noted) were set up with the following design. Unless otherwise stated, crosses in each experiment were conducted by crossing 10 female heterozygous Act5c-Ga4/CyO driver flies to 2 male homozygous transgene flies (both uninfected, unless otherwise noted: switching the gender for each genotype does not alter the effect, data not shown). The offspring of these crosses were used for all experiments, except where noted. As the Act5c-Gal4/CyO driver strain is heterozygous, when driver flies are crossed to homozygous transgene flies, half of the offspring express the gene (those that inherit the Act5c driver gene that produces the Gal4 transcription factor), while the other half do not (those that inherit the CyO chromosome, which does not produce Gal4). Therefore, expressing males, expressing females, non-expressing males, and non-expressing females are expected in equal proportions under Mendelian inheritance. These four genotypes can only be visibly assessed in adulthood. Visually, embryos cannot be distinguished (except when fixed for microscopy with the Y chromosome FISH probe, when sex can be distinguished), while larvae can only be differentiated by sex.
Alongside several experiments, including the cytology, sex ratios were measured concurrently. When flies were set up in the crosses described above, siblings were also set up in vials with CMY media. The protocol to measure sex ratios was then followed to obtain sex ratios side by side with these experiments.
The MTD driver was tested by crossing this homozygous driver strain to homozygous transgene flies in the same design as above. This crossing leads to gene expression in all offspring because the driver is homozygous. Females expressing the transgene in their ovaries (MTD leads to targeted gene expression in the germline, specifically by loading embryos with the product) were then crossed to WT flies. Offspring were then quantified to measure sex ratios.
Sex Ratio Measurements:
To assess the ability of transgene expression to alter sex ratios, twenty replicates of 10 uninfected, 4-7 day old female driver flies and 2 uninfected, 1-2 day old male transgene flies were set up in vials with CMY media (except
Hatch Rate:
To assess the hatch rate, 32 replicates each of 10 uninfected, 4-7 day old Act5c-Ga14/Cy0 females and 2 uninfected, 1-2 day old transgene males were set up in 8 oz, round bottom Drosophila polypropylene stock bottles. The hatch rate was then conducted as described previously10. After the embryo hatch counting, the sex of third instar larval offspring from each replicate was visually confirmed, and they were moved into their own CMY media vial where they developed to adulthood. The sex ratio of surviving adult offspring was then quantified. Any crosses with fewer than 50 embryos were not included in the analysis.
Hatch rates (for
Embryo Cytology:
Eight stock bottles were set up per genotype, each with 60 uninfected, 4-7 day old Act5c-Gal4/Cy0 females and 12 uninfected, 1-2 day old transgene or WT males. Grape juice agar plates, made as described previously10, with a small amount of baker's yeast (Red Star) were placed on each bottle opening and fixed on with tape. They were then placed with the grape plate down in a 25° C. incubator overnight (˜16 hr). Then, the grape plates were replaced with fresh plates and fresh yeast. The flies were then allowed to lay eggs in 1 h increments, replacing the previous plates with fresh ones each time. They were then allowed to sit at room temperature for 1 h (embryos 1-2 h old), 3 h (3-4 h old), or 16 h (16-17 hold). Once they had reached the desired point in development, the embryos were fixed and stained, using a slight modification of the protocol outlined by Cheng et al. 201639. Briefly, the embryos were dechorionated in 50% bleach and fixed for 15 minutes in a 1:1 4% paraformaldehyde: heptane mixture while shaking on a tabletop vortexer at about 150 rpm. The solution was discarded and the embryos were then devitellinized in a 1:1 heptane:methanol mixture by shaking vigorously for one minute. The solution was removed and the embryos were placed in fresh methanol and stored at 4° C. until the next steps were done, at least 16 h later. Then, the methanol was removed and the embryos were rehydrated in a series of methanol:water solutions, in the order of 9:1, then 1:1, then 1:9, each for 15 minutes while mixing on a Nutator. They were then treated with 10 mg/mL RNase A (Clontech Labs) by incubating them at 37° C. for 2-3 hr with enough RNase solution to cover the embryos. Once the RNase was removed, the embryos were washed three times for 5 min each in PBST (1×PBS, 0.1% Tween 20), while mixing on the Nutator. They were then re-fixed in 4% paraformaldehyde for 45 minutes with mixing and were then washed or incubated with several solutions with mixing on the nutator. First, they were washed three times in saline-sodium citrate/Tween 20 buffer (SSCT, 2×SSC buffer, 0.1% Tween 20) for 10 minutes each. They were then incubated with a series of SSCT/formamide solutions for 10 minutes each in the following order: 80% SSCT/20% formamide, 60% SSCT/40% formamide, 50% SSCT/50% formamide. Then fresh 50% SSCT/50% formamide was added and the embryos were incubated at 37° C. for 1 h. The solution was removed, and the embryos were then hybridized with the Y-chromosome FISH probe. This was done by mixing 36 μL FISH hybridization solution (1 g dextran sulfate, 1.5 mL 20×SSC, 5 mL formamide, to 15 mL with DNase-free water)40, 3 μL DNase-free water, and 1 μL 200 ng/μL Y-chromosome FISH probe (sequence 5′-AATACAATACAATACAATACAATACAATAC-3′ (SEQ ID NO:23) synthesized with Cy5 conjugated to the 5′end (IDT)) using the sequence published by Cheng et al. 201639. Hybridization was done in a thermocycler by denaturing at 92° C. for 3 min, followed by hybridizing at 37° C. overnight (˜16 h). Then, the embryos were again washed in a series of solutions on the nutator. They were done in the order of three 15 min 50% SSCT/50% formamide washes, one 10 min 60% SSCT/40% formamide wash, one 10 min 80% SSCT/20% formamide wash, and three 10 min SSCT washes. They were then mounted on glass slides with ProLong Diamond Antifade (Life Technologies) mounting media that contained 1 μg/mL propidium iodide (Sigma Aldrich).
Imaging was performed at the Vanderbilt University Cell Imaging Shared Resource (CISR) with a Zeiss LSM 510 META inverted confocal microscope. Image analysis and preparation was done with ImageJ software. Image brightness and contrast were adjusted for visibility, but adjustments were applied equally across each whole image.
Gene Expression:
Gene expression in embryos from
Confirmation of gene expression in adults from
Protein Conservation
Protein conservation was calculated with the Protein Residue Conservation Prediction Tool (68http://compbio.cs.princeton.edu/conservation/score.html). Amino acid sequences from
Statistical Analyses:
Statistical analyses were done using GraphPad Prism software (version 5) or GraphPad online tools. For comparisons among only two data categories, the two-tailed, non-parametric Mann-Whitney U test was used. For comparisons with more groups, a non-parametric Kruskal-Wallis one-way analysis of variance was used, followed by Dunn's test for multiple comparisons if significant. In cases of comparisons among groups where only a single measurement was taken per group (such as cytology experiments), a Chi-square test was used. Exact tests used and other important information is listed in the figure legends of each experiment.
REFERENCES CITED IN THIS EXAMPLE
- 1 LePage, D. & Bordenstein, S. R. Wolbachia: Can we save lives with a great pandemic? Trends Parasitol 29, 385-393, doi:10.1016/j.pt.2013.06.003 (2013).
- 2 Brucker, R. M. & Bordenstein, S. R. Speciation by symbiosis. Trends Ecol Evol 27, 443-451, doi:10.1016/j.tree.2012.03.011 (2012).
- 3 Hornett, E. A. et al. Evolution of male-killer suppression in a natural population. PLoS Biol 4, e283, doi:10.1371/journal.pbio.0040283 (2006).
- 4 Bordenstein, S. R., O'Hara, F. P. & Werren, J. H. Wolbachia-induced incompatibility precedes other hybrid incompatibilities in Nasonia. Nature 409, 707-710, doi:10.1038/35055543 (2001).
- 5 Jaenike, J., Dyer, K. A., Cornish, C. & Minhas, M. S. Asymmetrical reinforcement and Wolbachia infection in Drosophila. PLoS Biol 4, e325, doi:10.1371/journal.pbio.0040325 (2006).
- 6 Hoffmann, A. A. et al. Successful establishment of Wolbachia in Aedes populations to suppress dengue transmission. Nature 476, 454-457, doi:10.1038/nature10356 (2011).
- 7 Dutra, H. L. et al. Wolbachia Blocks Currently Circulating Zika Virus Isolates in Brazilian Aedes aegypti Mosquitoes. Cell Host Microbe 19, 771-774, doi:10.1016/j.chom.2016.04.021 (2016).
- 8 Bordenstein, S. R. & Bordenstein, S. R. Eukaryotic association module in phage WO genomes from Wolbachia. Nat Commun 7, 13155, doi:10.1038/ncomms13155 (2016).
- 9 Berec, L., Maxin, D. & Bernhauerova, V. Male-killing bacteria as agents of insect pest control. Journal of Applied Ecology (2016).
- 10 LePage, D. P. et al. Prophage WO genes recapitulate and enhance Wolbachia-induced cytoplasmic incompatibility. Nature 543, 243-247, doi:10.1038/nature21391 (2017).
- 11 Pinto, S. B. et al. Transcriptional regulation of Culex pipiens mosquitoes by Wolbachia influences cytoplasmic incompatibility. PLoS pathogens 9, e1003647, doi:10.1371/journal.ppat.1003647 (2013).
- 12 Sutton, E. R., Harris, S. R., Parkhill, J. & Sinkins, S. P. Comparative genome analysis of
Wolbachia strain wAu. BMC genomics 15, 928, doi:10.1186/1471-2164-15-928 (2014).
- 13 Beckmann, J. F., Ronau, J. A. & Hochstrasser, M. A Wolbachia deubiquitylating enzyme induces cytoplasmic incompatibility. Nature microbiology 2, 17007, doi:10.1038/nmicrobiol.2017.7 (2017).
- 14 Hurst, G. D., Johnson, A. P., Schulenburg, J. H. & Fuyama, Y. Male-killing Wolbachia in Drosophila: a temperature-sensitive trait with a threshold bacterial density. Genetics 156, 699-709 (2000).
- 15 Jaenike, J. Spontaneous emergence of a new Wolbachia phenotype. Evolution 61, 2244-2252, doi:10.1111/j.1558-5646.2007.00180.x (2007).
- 16 Sasaki, T., Kubo, T. & Ishikawa, H. Interspecific transfer of Wolbachia between two lepidopteran insects expressing cytoplasmic incompatibility: a Wolbachia variant naturally infecting Cadra cautella causes male killing in Ephestia kuehniella. Genetics 162, 1313-1319 (2002).
- 17 Metcalf, J. A., Jo, M., Bordenstein, S. R., Jaenike, J. & Bordenstein, S. R. Recent genome reduction of Wolbachia in Drosophila recens targets phage WO and narrows candidates for reproductive parasitism. PeerJ 2, e529, doi:10.7717/peerj.529 (2014).
- 18 Petrella, L. N., Smith-Leiker, T. & Cooley, L. The Ovhts polyprotein is cleaved to produce fusome and ring canal proteins required for Drosophila oogenesis. Development 134, 703-712, doi:10.1242/dev.02766 (2007).
- 19 Landmann, F., Orsi, G. A., Loppin, B. & Sullivan, W. Wolbachia-mediated cytoplasmic incompatibility is associated with impaired histone deposition in the male pronucleus. PLoS pathogens 5, e1000343, doi:10.1371/journal.ppat.1000343 (2009).
- 20 Riparbelli, M. G., Giordano, R., Ueyama, M. & Callaini, G. Wolbachia-mediated male killing is associated with defective chromatin remodeling. PloS one 7, e30045, doi:10.1371/journal.pone.0030045 (2012).
- 21 Dyer, K. A. & Jaenike, J. Evolutionarily stable infection by a male-killing endosymbiont in Drosophila innubila: molecular evidence from the host and parasite genomes. Genetics 168, 1443-1455, doi:10.1534/genetics.104.027854 (2004).
- 22 Carson, H. L. A female-producing strain of D. borealis Patterson. Drosoph. Inf. Serv 30, 109-110 (1956).
- 23 Hornett, E. A. et al. You can't keep a good parasite down: evolution of a male-killer suppressor uncovers cytoplasmic incompatibility. Evolution 62, 1258-1263, doi:10.1111/j.1558-5646.2008.00353.x (2008).
- 24 Kelley, L. A., Mezulis, S., Yates, C. M., Wass, M. N. & Sternberg, M. J. The Phyre2 web portal for protein modeling, prediction and analysis. Nature protocols 10, 845-858, doi:10.1038/nprot.2015.053 (2015).
- 25 Kim, M. et al. Noncanonical DNA-binding mode of repressor and its disassembly by antirepressor. Proceedings of the National Academy of Sciences of the United States of America 113, E2480-2488, doi:10.1073/pnas.1602618113 (2016).
- 26 Luscombe, N. M., Austin, S. E., Berman, H. M. & Thornton, J. M. An overview of the structures of protein-DNA complexes. Genome biology 1, Reviews001, doi:10.1186/gb-2000-1-1-reviews001 (2000).
- 27 Engelstadter, J. & Hurst, G. D. The impact of male-killing bacteria on host evolutionary processes. Genetics 175, 245-254, doi:10.1534/genetics.106.060921 (2007).
- 28 Telschow, A., Hammerstein, P. & Werren, J. H. The effect of Wolbachia versus genetic incompatibilities on reinforcement and speciation. Evolution 59, 1607-1619 (2005).
29 Hurst, G. D. & McVean, G. A. T. Parasitic male-killing bacteria and the evolution of clutch size. Ecological entomology 23, 350-353 (1998).
- 30 Majerus, T. M. & Majerus, M. E. Intergenomic arms races: detection of a nuclear rescue gene of male-killing in a ladybird. PLoS pathogens 6, e1000987, doi:10.1371/journal.ppat.1000987 (2010).
- 31 Jiggins, F. M., Hurst, G. D. & Majerus, M. E. Sex-ratio-distorting Wolbachia causes sex-role reversal in its butterfly host. Proc Biol Sci 267, 69-73, doi:10.1098/rspb.2000.0968 (2000).
- 32 Hurst, G. D. & Jiggins, F. M. Male-killing bacteria in insects: mechanisms, incidence, and implications. Emerg Infect Dis 6, 329-336, doi:10.3201/eid0604.000402 (2000).
- 33 Baym, M. et al. Inexpensive multiplexed library preparation for megabase-sized genomes. PloS one 10, e0128036, doi:10.1371/journal.pone.0128036 (2015).
- 34 Bankevich, A. et al. SPAdes: a new genome assembly algorithm and its applications to single-cell sequencing. Journal of computational biology: a journal of computational molecular cell biology 19, 455-477, doi:10.1089/cmb.2012.0021 (2012).
- 35 Longdon, B., Fabian, D. K., Hurst, G. D. & Jiggins, F. M. Male-killing Wolbachia do not protect Drosophila bifasciata against viral infection. BMC microbiology 12 Suppl 1, S8, doi:10.1186/1471-2180-12-s1-s8 (2012).
- 36 Sheeley, S. L. & McAllister, B. F. Mobile male-killer: similar Wolbachia strains kill males of divergent Drosophila hosts. Heredity 102, 286-292, doi:10.1038/hdy.2008.126 (2009).
- 37 Ellegaard, K. M., Klasson, L., Naslund, K., Bourtzis, K. & Andersson, S. G. Comparative genomics of Wolbachia and the bacterial species concept. PLoS genetics 9, e1003381, doi:10.1371/journal.pgen.1003381 (2013).
- 38 Boetzer, M. & Pirovano, W. Toward almost closed genomes with GapFiller. Genome biology 13, R56, doi:10.1186/gb-2012-13-6-r56 (2012).
- 39 Cheng, B., Kuppanda, N., Aldrich, J. C., Akbari, O. S. & Ferree, P. M. Male-Killing Spiroplasma Alters Behavior of the Dosage Compensation Complex during Drosophila melanogaster Embryogenesis. Curr Biol 26, 1339-1345, doi: 10.1016/j .cub.2016.03.050 (2016).
- 40 Sullivan, W., Ashburner, M. & Hawley, R. S. Drosophila protocols. (Cold Spring Harbor Laboratory Press, 2000).
Tables
Unless defined otherwise, all technical and scientific terms used herein have the same meanings as commonly understood by one of skill in the art to which the disclosed invention belongs. Publications cited herein and the materials for which they are cited are specifically incorporated by reference.
Those skilled in the art will appreciate that numerous changes and modifications can be made to the preferred embodiments of the invention and that such changes and modifications can be made without departing from the spirit of the invention. It is, therefore, intended that the appended claims cover all such equivalent variations as fall within the true spirit and scope of the invention.
Claims
1. A genetically modified arthropod, said arthropod comprising:
- a gene encoding a male arthropod killing factor; and
- a promoter operably linked to the gene encoding the male arthropod killing factor;
- wherein the expression of the gene encoding the male arthropod killing factor in arthropod embryos causes a reduction in viable surviving male offspring in comparison to arthropod embryos not expressing the gene encoding the male arthropod killing factor; and
- wherein the male arthropod killing factor comprises the amino acid sequence SEQ ID NO:1, or an amino acid sequence at least 60% identical to SEQ ID NO:1.
2. The arthropod of claim 1, wherein the gene encoding the male arthropod killing factor is from a bacterium, a prophage, or a phage.
3. The arthropod of claim 2, wherein the gene encoding the male arthropod killing factor is from Wolbachia.
4. The arthropod of claim 3, wherein the male arthropod killing factor is wmk (WD0626).
5. The arthropod of claim 1, wherein the reduction in viable male offspring is greater than 10%.
6. The arthropod of claim 1, wherein the arthropod is an insect.
7. The arthropod of claim 6, wherein the insect is selected from the genera consisting of Aedes, Culex and Anopheles.
8. The arthropod of claim 7, wherein the insect is selected from the group consisting of Aedes albopictus, Aedes aegypti and Aedes polynesiensis.
9. The arthropod of claim 6, wherein the insect is Drosophila suzukii.
10. A method for controlling a population of target arthropods, comprising:
- providing a gene encoding a male arthropod killing factor, and a promoter operably linked to the gene encoding the male arthropod killing factor; and
- wherein the male arthropod killing factor comprises the amino acid sequence SEQ ID NO:1, or an amino acid sequence at least 60% identical to SEQ ID NO:1;
- transforming a population of arthropods with the gene encoding the male arthropod killing factor and the promoter operably linked to the gene encoding the male arthropod-killing factor; and
- releasing the population of arthropods amongst a population of target arthropods, wherein the release of the arthropods reduces the population of target arthropods.
11. The method of claim 10, wherein the gene encoding the male arthropod killing factor is from a bacterium, a prophage, or a phage.
12. The method of claim 11, wherein the gene encoding the male arthropod killing factor is from Wolbachia.
13. The method of claim 12, wherein the gene encoding the male arthropod killing factor is wmk (WD0626).
14. The method of claim 10, wherein the male arthropod killing factor comprises the amino acid sequence SEQ ID NO: 1.
15. The method of claim 10, wherein the reduction in viable male offspring is greater than 10%.
16. The method of claim 10, further comprising providing an additional method of arthropod control.
17. The method of claim 16, wherein the additional method of arthropod control is a sterile insect technique (SIT).
18. The method of claim 16, wherein the additional method of arthropod control is an incompatible insect technique (ITT).
19. A method for controlling a population of target arthropods, comprising:
- providing a gene encoding a male arthropod killing factor, and a promoter operably linked to the gene encoding the male arthropod killing factor; and
- wherein the male arthropod killing factor comprises the amino acid sequence SEQ ID NO:1, or an amino acid sequence at least 60% identical to SEQ ID NO:1;
- genetically transforming a bacteria, phage, or prophage with the gene encoding the male arthropod killing factor operably linked to the promoter;
- transinfecting a population of arthropods with the bacteria, phage, or prophage; and
- releasing the population of arthropods amongst a population of target arthropods, wherein the release of the arthropods reduces the population of target arthropods.
| 7868222 | January 11, 2011 | Dobson |
| 9090911 | July 28, 2015 | O'Neill et al. |
| 9125388 | September 8, 2015 | Alphey et al. |
| 20130259846 | October 3, 2013 | Dobson |
| 20170188559 | July 6, 2017 | Koukidou et al. |
| 2017/181043 | October 2017 | WO |
| 2017/185041 | October 2017 | WO |
| 2017/214476 | December 2017 | WO |
- Arai et al. (Frontiers in Microbiology, 2022, 13, 1-15).
- Aliota, Matthew T., et al. “The w Mel strain of Wolbachia reduces transmission of chikungunya virus in Aedes aegypti.” PLOS neglected tropical diseases 10.4 (2016): e0004677.
- Altschul, S. F., Gish, W., Miller, W., Myers, E. W., & Lipman, D. J. (1990). Basic local alignment search tool. Journal of molecular biology, 215(3), 403-410.
- Altschul, S. F., Madden, T. L., Schäffer, A. A., Zhang, J., Zhang, Z., Miller, W., & Lipman, D. J. (1997). Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic acids research, 25(17), 3389-3402.
- Bankevich, A. et al. SPAdes: a new genome assembly algorithm and its applications to single-cell sequencing. Journal of computational biology: a journal of computational molecular cell biology 19, 455-477, doi: 10.1089/cmb.2012.0021 (2012).
- Baumhover, A. H., et al. “Field observations on the effects of releasing sterile screw-worms in Florida.” Journal of Economic Entomology 52.6 (1959): 1202-1206.
- Baumhover, Alfred H. “Eradication of the screwworm fly: an agent of myiasis.” JAMA 196.3 (1966): 240-248.
- Baumhover, et al., Screw-Worm Control Through Release of Sterilized Flies, Journal of Economic Entomology, vol. 48, Issue 4, Aug. 1, 1955, pp. 462-466, https://doi.org/10.1093/jee/48.4.462.
- Baym, M. et al. Inexpensive multiplexed library preparation for megabase-sized genomes. PloS one 10, e0128036, doi:10.1371/journal.pone.0128036 (2015).
- Beaucage, S. L., & Caruthers, M. H. (1981). Deoxynucleoside phosphoramidites—a new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Letters, 22(20), 1859-1862.
- Beckmann, J. F., Ronau, J. A. & Hochstrasser, M. A Wolbachia deubiquitylating enzyme induces cytoplasmic incompatibility. Nature microbiology 2, 17007, doi:10.1038/nmicrobiol.2017.7 (2017).
- Benedict, Mark Q., and Alan S. Robinson. “The first releases of transgenic mosquitoes: an argument for the sterile insect technique.” Trends in parasitology 19.8 (2003): 349-355.
- Berec, L., Maxin, D. & Bernhauerova, V. Male-killing bacteria as agents of insect pest control. Journal of Applied Ecology (2016).
- Boetzer, M. & Pirovano, W. Toward almost closed genomes with GapFiller. Genome biology 13, R56, doi:10.1186/gb-2012-13-6-r56 (2012).
- Bordenstein, S. R. & Bordenstein, S. R. Eukaryotic association module in phage WO genomes from Wolbachia. Nat Commun 7, 13155, doi:10.1038/ncomms13155 (2016).
- Bordenstein, S. R., O'Hara, F. P. & Werren, J. H. Wolbachia-induced incompatibility precedes other hybrid incompatibilities in Nasonia. Nature 409, 707-710, doi:10.1038/35055543 (2001).
- Brelsfoard, Corey L., and Stephen L. Dobson. “Wolbachia-based strategies to control insect pests and disease vectors.” Asia Pac. J. Mol. Biol. Biotechnol 17.3 (2009): 55-63.
- Brucker, R. M. & Bordenstein, S. R. Speciation by symbiosis. Trends Ecol Evol 27, 443-451, doi:10.1016/j.tree.2012.03.011 (2012).
- Bushland, B. C., A. W. Lindquist, and E. F. Kipling. “Eradication of Screw-Worms through Release of Sterilized Males.” Science (Washington) 122 (1955): 287-8.
- Bushland, R. C., and D. E. Hopkins. “Experiments with screw-worm flies sterilized by X-rays.” Journal of Economic Entomology 44.5 (1951).
- Carson, H. L. A female-producing strain of D. borealis Patterson. Drosoph. Inf. Serv 30, 109-110 (1956).
- Cheng, B., Kuppanda, N., Aldrich, J. C., Akbari, O. S. & Ferree, P. M. Male-Killing Spiroplasma Alters Behavior of the Dosage Compensation Complex during Drosophila melanogaster Embryogenesis. Curr Biol 26, 1339-1345, doi: 10.1016/j.cub.2016.03.050 (2016).
- Dame, David A., et al. “Historical applications of induced sterilisation in field populations of mosquitoes.” Malaria journal 8.2 (2009): 1-10.
- Dutra, H. L. et al. Wolbachia Blocks Currently Circulating Zika Virus Isolates in Brazilian Aedes aegypti Mosquitoes. Cell Host Microbe 19, 771-774, doi:10.1016/j.chom.2016.04.021 (2016).
- Dyer, K. A. & Jaenike, J. Evolutionarily stable infection by a male-killing endosymbiont in Drosophila innubila: molecular evidence from the host and parasite genomes. Genetics 168, 1443-1455, doi:10.1534/genetics.104.027854 (2004).
- Ellegaard, K. M., Klasson, L., Naslund, K., Bourtzis, K. & Andersson, S. G. Comparative genomics of Wolbachia and the bacterial species concept. PLoS genetics 9, e1003381, doi: 10.1371/journal.pgen.1003381 (2013).
- Engelstadter, J. & Hurst, G. D. The impact of male-killing bacteria on host evolutionary processes. Genetics 175, 245-254, doi:10.1534/genetics.106.060921 (2007).
- Henikoff, S., & Henikoff, J. G. (1992). Amino acid substitution matrices from protein blocks. Proceedings of the National Academy of Sciences, 89(22), 10915-10919.
- Hoffmann, A. A. et al. Successful establishment of Wolbachia in Aedes populations to suppress dengue transmission. Nature 476, 454-457, doi:10.1038/nature10356 (2011).
- Hornett, E. A. et al. Evolution of male-killer suppression in a natural population. PLoS Biol 4, e283, doi:10.1371/journal.pbio.0040283 (2006).
- Hornett, E. A. et al. You can't keep a good parasite down: evolution of a male-killer suppressor uncovers cytoplasmic incompatibility. Evolution 62, 1258-1263, doi:10.1111/j.1558-5646.2008.00353.x (2008).
- Hughes, Grant L., et al. “Native microbiome impedes vertical transmission of Wolbachia in Anopheles mosquitoes.” Proceedings of the National Academy of Sciences 111.34 (2014): 12498-12503.
- Hughes, Grant L., et al. “Wolbachia infections are virulent and inhibit the human malaria parasite Plasmodium falciparum in Anopheles gambiae.” PLoS Pathog 7.5 (2011): e1002043.
- Hurst, G. D. & Jiggins, F. M. Male-killing bacteria in insects: mechanisms, incidence, and implications. Emerg Infect Dis 6, 329-336, doi:10.3201/eid0604.000402 (2000).
- Hurst, G. D. & McVean, G. A. T. Parasitic male-killing bacteria and the evolution of clutch size. Ecological entomology 23, 350-353 (1998).
- Hurst, G. D., Johnson, A. P., Schulenburg, J. H. & Fuyama, Y. Male-killing Wolbachia in Drosophila: a temperature-sensitive trait with a threshold bacterial density. Genetics 156, 699-709 (2000).
- Jaenike, J. Spontaneous emergence of a new Wolbachia phenotype. Evolution 61, 2244-2252, doi:10.1111/j.1558-5646.2007.00180.x (2007).
- Jaenike, J., Dyer, K. A., Cornish, C. & Minhas, M. S. Asymmetrical reinforcement and Wolbachia infection in Drosophila. PLoS Biol 4, e325, doi:10.1371/journal.pbio.0040325 (2006).
- Jiggins, F. M., Hurst, G. D. & Majerus, M. E. Sex-ratio-distorting Wolbachia causes sex-role reversal in its butterfly host. Proc Biol Sci 267, 69-73, doi:10.1098/rspb.2000.0968 (2000).
- Karlin, S., & Altschul, S. F. (1993). Applications and statistics for multiple high-scoring segments in molecular sequences. Proceedings of the National Academy of Sciences, 90(12), 5873-5877.
- Kelley, L. A., Mezulis, S., Yates, C. M., Wass, M. N. & Sternberg, M. J. The Phyre2 web portal for protein modeling, prediction and analysis. Nature protocols 10, 845-858, doi: 10.1038/nprot.2015.053 (2015).
- Kim, M. et al. Noncanonical DNA-binding mode of repressor and its disassembly by antirepressor. Proceedings of the National Academy of Sciences of the United States of America 113, E2480-2488, doi:10.1073/pnas.1602618113 (2016).
- Knipling, E. F. “Possibilities of insect control or eradication through the use of sexually sterile males.” Journal of Economic Entomology 48.4 (1955): 459-462.
- Lacroix, Renaud, et al. “Open field release of genetically engineered sterile male Aedes aegypti in Malaysia.” PloS one 7.8 (2012): e42771.
- Landmann, F., Orsi, G. A., Loppin, B. & Sullivan, W. Wolbachia-mediated cytoplasmic incompatibility is associated with impaired histone deposition in the male pronucleus. PLoS pathogens 5, e1000343, doi:10.1371/journal.ppat.1000343 (2009).
- Laven, H. “Eradication of Culex pipiens fatigans through cytoplasmic incompatibility.” Nature 216.5113 (1967): 383-384.
- Lees, Rosemary Susan, et al. “Back to the future: the sterile insect technique against mosquito disease vectors.” Current Opinion in Insect Science 10 (2015): 156-162.
- LePage, D. & Bordenstein, S. R. Wolbachia: Can we save lives with a great pandemic? Trends Parasitol 29, 385-393, doi:10.1016/j.pt.2013.06.003 (2013).
- LePage, D. P. et al. Prophage WO genes recapitulate and enhance Wolbachia-induced cytoplasmic incompatibility. Nature 543, 243-247, doi:10.1038/nature21391 (2017).
- Lindquist, D. A., M. Abusowa, and M. J. R. Hall. “The New World screwworm fly in Libya: a review of its introduction and eradication.” Medical and Veterinary Entomology 6.1 (1992): 2-8.
- Longdon, B., Fabian, D. K., Hurst, G. D. & Jiggins, F. M. Male-killing Wolbachia do not protect Drosophila bifasciata against viral infection. BMC microbiology 12 Suppl 1, S8, doi:10.1186/1471-2180-12-s1-s8 (2012).
- Luscombe, N. M., Austin, S. E., Berman, H. M. & Thornton, J. M. An overview of the structures of protein-DNA complexes. Genome biology 1, Reviews001, doi:10.1186/gb-2000-1-1-reviews001 (2000).
- Mains, James W., et al. “Female adult Aedes albopictus suppression by Wolbachia-infected male mosquitoes.” Scientific reports 6 (2016): 33846.
- Majerus, T. M. & Majerus, M. E. Intergenomic arms races: detection of a nuclear rescue gene of male-killing in a ladybird. PLoS pathogens 6, e1000987, doi:10.1371/journal.ppat.1000987 (2010).
- Matteucci, M. D., & Caruthers, M. H. (1981). Synthesis of deoxyoligonucleotides on a polymer support. Journal of the American Chemical Society, 103(11), 3185-3191.
- Metcalf, J. A., Jo, M., Bordenstein, S. R., Jaenike, J. & Bordenstein, S. R. Recent genome reduction of Wolbachia in Drosophila recens targets phage WO and narrows candidates for reproductive parasitism. PeerJ 2, e529, doi:10.7717/peerj.529 (2014).
- Petrella, L. N., Smith-Leiker, T. & Cooley, L. The Ovhts polyprotein is cleaved to produce fusome and ring canal proteins required for Drosophila oogenesis. Development 134, 703-712, doi:10.1242/dev.02766 (2007).
- Pinto, S. B. et al. Transcriptional regulation of Culex pipiens mosquitoes by Wolbachia influences cytoplasmic incompatibility. PLoS pathogens 9, e1003647, doi:10.1371/journal.ppat.1003647 (2013).
- Riparbelli, M. G., Giordano, R., Ueyama, M. & Callaini, G. Wolbachia-mediated male killing is associated with defective chromatin remodeling. PloS one 7, e30045, doi:10.1371/journal.pone.0030045 (2012).
- Sasaki, T., Kubo, T. & Ishikawa, H. Interspecific transfer of Wolbachia between two lepidopteran insects expressing cytoplasmic incompatibility: a Wolbachia variant naturally infecting Cadra cautella causes male killing in Ephestia kuehniella. Genetics 162, 1313-1319 (2002).
- Sheeley, S. L. & McAllister, B. F. Mobile male-killer: similar Wolbachia strains kill males of divergent Drosophila hosts. Heredity 102, 286-292, doi:10.1038/hdy.2008.126 (2009).
- Sinkins, Steven P. “Wolbachia and cytoplasmic incompatibility in mosquitoes.” Insect biochemistry and molecular biology 34.7 (2004): 723-729.
- Sinkins, Steven P., and Fred Gould. “Gene drive systems for insect disease vectors.” Nature Reviews Genetics 7.6 (2006): 427-435.
- Sutton, E. R., Harris, S. R., Parkhill, J. & Sinkins, S. P. Comparative genome analysis of Wolbachia strain wAu. BMC genomics 15, 928, doi:10.1186/1471-2164-15-928 (2014).
- Telschow, A., Hammerstein, P. & Werren, J. H. The effect of Wolbachia versus genetic incompatibilities on reinforcement and speciation. Evolution 59, 1607-1619 (2005).
- Walker, T. J. P. H., et al. “The w Mel Wolbachia strain blocks dengue and invades caged Aedes aegypti populations.” Nature 476.7361 (2011): 450-453.
- International Search Report and Written Opinion dated Jul. 5, 2019, from International Application No. PCT/US2019/025936, 13 pages.
- Sutton et al. “Comparative genome analysis of Wolbachia strain wAu”, BMC Genomics 2014, 15:928.
- Riparbelli et al. “Wolbachia-Mediated Male Killing is Associated with Defective Chromatin Remodeling”, PLOS One, Jan. 2012, vol. 7, issue 1, 14 pages.
Type: Grant
Filed: Apr 5, 2019
Date of Patent: Apr 30, 2024
Patent Publication Number: 20210000092
Assignee: VANDERBILT UNIVERSITY (Nashville, TN)
Inventors: Seth Bordenstein (Nashville, TN), Jessamyn Perlmutter (Nashville, TN)
Primary Examiner: Amy H Bowman
Application Number: 16/982,708
