Patent Application
20200045945
The present invention relates to genetically edited swine which produce CD163 protein in which the scavenger receptor cysteine-rich 5 (SRCR5) domain has been deleted. Such swine have been found to be healthy and do not exhibit negative properties, and are resistant to PRRSV infection. Moreover, the CD163 protein without the SRCR5 retains the ability to function as a haemoglobin-haptoglobin scavenger. The invention also relates to methods of producing such swine.
Porcine Reproductive and Respiratory Syndrome Virus (PRRSV) is a virus that causes a disease of pigs, called Porcine Reproductive and Respiratory Syndrome (PRRS).
This economically important disease, which is endemic in many pig producing countries, causes reproductive failure in breeding stock and respiratory tract illness in young pigs. Initially referred to as “mystery swine disease” and “mystery reproductive syndrome,” it was first reported in 1987 in North America and Central Europe. It is estimated that the disease costs the United States swine industry around $650 million annually.
PRRSV enters macrophages via a set of macrophage cell surface markers: CD169 and CD163. The role of CD169/sialoadhesin was discovered by the group of Hans Nauwynck in Ghent. The role of CD163 was discovered by scientists working with Pfizer (Calvert et al. 2007). Calvert et al. (2007) demonstrated that transfection of any non-susceptible cells with CD163 can render the cells susceptible to PRRSV. That has allowed for the generation of vaccine strains without the need of using Marc-145 cells.
Van Gorp et al. (“Susceptible cell lines for the production of porcine reproductive and respiratory syndrome virus by stable transfection of sialoadhesin and CD163”, BMC Biotechnology 2010, 10:48) have demonstrated that the domains 5 to 9 of the CD163 protein are important for the PRRSV entry into non-susceptible cells and highlighted that domain 5 may be critical.
Das et al. (“The Minor Envelope Glycoproteins GP2a and GP4 of Porcine Reproductive and Respiratory Syndrome Virus Interact with the Receptor CD163”, JOURNAL OF VIROLOGY, February 2010, p. 1731-1740) have demonstrated that that the PRRSV glycoprotein GP2A and GP4 interact physically with CD163.
US 20050271685 held by Pfizer (Zoetis) suggests that the use of CD163 molecule can make cells susceptible to PRRSV and ASFV.
WO 2012/158828 describes PRRS resistant animals in which the SIGLEC1 and/or CD163 genes have been inactivated. CD163, however, has roles in normal physiological activities. It is therefore undesirable to inactive this gene as it may have undesirable and unforeseeable knock-on effects on the animal.
There remains a need for improvements in the prevention and treatment of PRRSV.
The present inventors have succeeded in generating genetically edited swine which produces CD163 in which the scavenger receptor cysteine-rich 5 (SRCR5) domain (also known as CD163 domain 5) has been deleted. Swine produced by the inventors have been found to be healthy and do not exhibit negative properties. Experiments conducted by the inventors have shown that the swine demonstrate resistance to PRRSV infection. CD163 expressed in the edited swine also demonstrates retention of the ability to function as a haemoglobin-haptoglobin scavenger.
According to a first aspect of the present invention, there is provided a genetically edited swine, the swine comprising an edited genome wherein the edit results in the deletion of SRCR5 domain from CD163 protein produced by the swine. In other words, the genetically edited swine produces a modified form of the CD163 protein in which SRCR5 (also referred to in context as domain 5) is absent.
Preferably the swine is a pig (Sus scrofa), and most preferably a domestic pig (Sus scrofa domesticus or Sus domesticus).
Suitably the swine comprises an edited genome wherein the edit results in the deletion of SRCR5 from CD163 protein produced by the animal, and wherein all of the other CD163 domains are present and their amino acid sequences are unaltered. Accordingly, the swine suitably produces CD163 in which SRCR5 is absent, but SRCR domains 1 to 4 and 6 to 9 are unaltered, as are the transmembrane segment and cytoplasmic domain. The present inventors have found, surprisingly, that a CD163 protein in which SRCR5 has been deleted can retain its physiological function as a hemoglobin-haptoglobin scavenger, but generates high levels of resistance to infection by PRRSV in cells bearing the modified CD163 protein.
Accordingly, in certain embodiments of the present invention the CD163 protein expressed from the edited genome preferably remains substantially functional. ‘Substantially functional’ in this context refers to the protein retaining physiological functions that are not dependent on the SRCR5 domain. Suitably the modified CD163 protein is substantially functional, in that it is able to function as a haemoglobin-haptoglobin scavenger. The ability of a CD163 protein to function as a haemoglobin-haptoglobin scavenger can readily be assessed according to the methodology described herein, i.e. based upon the ability of peripheral blood monocyte-derived macrophages from edited swine to scavenge haemoglobin-haptoglobin. The ability of the CD163 protein to function as a haemoglobin-haptoglobin scavenger is indicative that the CD163 protein is correctly folded and functional despite the deletion of the SRCR5 domain.
SRCR5 of CD163 has the following amino acid sequence:
Accordingly, the modified CD163 protein produced by the edited swine suitably lacks the abovementioned amino acid sequence, i.e. SEQ ID NO:2. Suitably the CD163 protein produced by the edited swine has no further changes to the wild type amino acid sequence.
The swine is preferably homozygous or biallelic for the genome edit that results in the deletion of SRCR5 from CD163 produced by the animal. ‘Homozygous’ in this context means that the swine comprises the same edit within the CD163 gene on both chromosomes, i.e. it has identical alleles on both chromosomes. ‘Biallelic’ in this context means the swine has different edits on each chromosome, but wherein both of the edits result in a desirable edit to CD163, i.e. which results in the deletion of SRCR5 from CD163 protein produced by the animal.
Preferably all cells of the animal comprise the edited genome. In some cases, however, the animal can exhibit mosaicism, with some cells comprising the edited genome, and other cells not comprising the edited genome. PRRSV infects macrophages, and thus provided macrophages, and their progenitor cells, do not express CD163 which comprises SRCR5, the animals will be resistant to PRRSV infection.
It is generally preferred that the swine does not produce any CD163 which comprises SRCR5, i.e. all cells of the animal are homozygous or biallelic for the genome edit that results in the deletion of SRCR5 from CD163 produced by the swine.
It will be apparent to the skilled person that a genetically edited swine of the present invention can be a swine that has been directly subjected to a gene editing methodology as described herein, or a descendent of such a swine that retains the edited genome. Indeed, a swine that has been subjected to a gene editing methodology will in some cases be heterozygous, and will be bred to arrive at a homozygous or biallelic descendent.
Suitably the genome is edited such that the sequence which codes for SRCR5 is absent from the mRNA (preferably the mature mRNA) produced from the edited CD163 gene. This can be achieved by an edit that deletes exon 7, which encodes the SRCR5 domain of the CD163 protein, from the CD163 gene, or by an edit that results in the removal of the RNA sequence encoded by exon 7 from the transcript from the edited CD163 gene, e.g. as a result of splicing during the formation of mRNA.
Accordingly, in certain embodiments of the invention exon 7 of the CD163 gene is deleted. Deletion of exon 7 of the CD163 gene will of course result in the deletion of SRCR5 from the encoded CD163 protein.
In certain embodiments of the invention the splice acceptor site located at the 5′ of exon 7 is inactivated. Inactivation of the splice acceptor site at the 5′ end of exon 7 results in exon 7 being spliced out of the mRNA produced form the edited CD163 gene, thus deleting SRCR5 from the CD163 protein that is obtained from the mRNA when it is translated.
In embodiments of the invention where the swine comprises an edited genome in which exon 7 of the CD163 gene has been deleted, this can be achieved in various ways. For example, the deletion can be limited to exon 7, or the deletions can extend beyond exon 7 into flanking intronic regions (i.e. into introns 6 and 7). It is typically preferred that the entirety of exon 7 is deleted.
Suitably the edited genome is edited such that exon 7 has been deleted, but there are no other changes to other coding regions of the CD163 regions. In particular, it is typically preferred that no other exons of CD163 have been altered compared to the unedited genome. Accordingly, exons 1 to 6 and 8 to 16 are preferably unaltered.
In some embodiments, exon 7 and portions of introns 6 and 7, which flank exon 7, are deleted, but there are no other alterations in the remaining regions of the CD163 gene.
Exon 7 spans from position 23392 to position 23706 with reference to SEQ ID NO:1. Accordingly, this region is suitably deleted in the edited swine genome.
It should be noted that, while positions or regions in the CD163 gene are described herein with reference to SEQ ID NO: 1, there will be variations in sequence of the CD163 between different individual swine (e.g. where single nucleotide polymorphisms (SNPs) or other polymorphisms occur), and thus individual swine may comprise a CD163 sequence that is slightly different to SEQ ID NO:1. References to positions or regions made with reference to SEQ ID NO: 1 are not meant to be strictly limiting, but should be construed as representative of the corresponding position in the CD163 gene of swine having any such sequence variation. The person skilled in the art could readily identify corresponding positions or regions in a CD63 gene comprising sequence variations using convention sequence alignment techniques, e.g. BLAST.
Suitably the edited genome is edited such that the splice site donor sequence in intron 6 (i.e. located at the junction of exon 6 and intron 6) and the splice site acceptor site in intron 7 (i.e. located at the junction of intron 7 and exon 8) are unaltered and remain functional. This facilitates correct splicing of the transcript produced from the edited CD163 gene. Accordingly, in embodiments of the present invention the sequences in the regions extending from position 10451 to position 10465, and from position 23783 to position 23824, with reference to SEQ ID NO: 1, are unaltered.
Suitably the genome is edited such that at least a portion of the region of the CD163 gene extending from position 10466 to 23782 with reference to SEQ ID NO:1 is deleted, wherein the portion comprises exon 7. Position 10466 lies immediately 3′ of the predicted splice donor site of intron 6 (i.e. at the 3′ end of exon 6). Position 23782 lies immediately 5′ of the predicted splice acceptor site of intron 7 (i.e. at the 5′ end of exon 8). The region can of course be smaller, provided that it comprises exon 7.
Suitably the genome is edited such that regions from positions 1 to position 10465 and from position 23783 or 23754 to position 32908, with reference to SEQ ID NO:1, are unaltered.
In certain embodiments of the present invention exon 7 is deleted along with up to 5000 bases, suitably up to 2000 bases, suitably up to 1000 bases, suitably up to 500 bases, suitably up to 300 bases or suitably up to 100 bases extending 5′ of the 5′ end of exon 7.
In certain embodiments of the present invention exon 7 is deleted along with up to 75 bases extending 3′ of the 3′ end of exon 7. This region extends from the 3′ end of exon 7 up to the predicted splice acceptor site at the 5′ end of exon 8. Suitably exon 7 is deleted along with up to 50 bases extending 3′ of the 3′ end of exon 7.
In one embodiment, the edited genome comprises a deletion of the region extending from approximately position 23060 to approximately position 23760, for example from position 23064 or 23065 to position 23753 or 23754, suitably 20365 to position 23753, with reference to SEQ ID NO:1.
In another embodiment, the edited genome comprises a deletion of the region extending from approximately position 23260 to approximately position 23760, for example from position 23267 or 23268 to position 237543 or 23754, suitably position 23268 to position 23753, with reference to SEQ ID NO:1.
In another embodiment, the edited genome comprises a deletion of the region extending from approximately position 23370 to approximately position 23760, for example from position 23373 or 23374 to position 237543 or 23754, suitably position 23374 to position 23753, with reference to SEQ ID NO:1.
In some embodiments of the invention the edited genome can comprise an inserted sequence not normally found at the relevant position (i.e. a heterologous inserted sequence). For example, when a section of the CD163 gene comprising exon 7 has been deleted, an inserted sequence can be present in the location in where the deletion occurred. Such insertions are a relatively common artefact of deletion of a sequence through gene editing. Such an insertion is typically inconsequential in the present context, and the inserted sequence is typically spliced out of the transcript produced from the gene. Accordingly, the inserted sequence typically does not result in any particular effect. The inserted sequence is generally not a sequence from the CD163 gene or any homologue or other related sequence. It is typically preferred that such a heterologous inserted sequence is not present in the edited genome.
In one particularly preferred embodiment the edited genome comprises a deletion of the region extending from position 23268 to position 23753, and wherein there is no insertion of a sequence at the location of the deletion. In such an embodiment, the edited genome of the swine at the former locus of the deleted exon 7 has the following sequence ATTGTCTCCAGGGAAGGACAGGGAGGTCTAGAATCGGCTAAGCCCAC∥GTAGGGTTAGGT AGTCA—SEQ ID NO:36 (wherein II represents the adjoining of the two cut sites that may be used to excise the region).
In certain embodiments of the invention, the swine comprises an edited genome in which the splice acceptor site in intron 6, i.e. located at the 5′ end of exon 7, of the CD163 gene has been inactivated. As mentioned above, inactivation of splice acceptor site at the 5′ end of exon 7 results in exon 7 being spliced out of the mRNA produced form the edited CD163 gene, thus deleting SRCR5 from the CD163 protein translated from the mRNA.
The predicted splice acceptor site in intron 6 extends from position 23378 to position 23416, with reference to SEQ ID NO:1. Accordingly, this sequence is suitably edited to inactivate the splice acceptor site.
The splice acceptor site can be partially or entirely deleted, or its sequence altered in any other suitable way so that it is no longer functional. Accordingly, in one embodiment the splice acceptor site is deleted. In another embodiment a sequence is inserted into the splice acceptor site that results in its inactivation. In another embodiment the splice acceptor site is modified such that it is inactivated, e.g. though a homology directed repair (HDR) mediated introgression event.
In one embodiment the sequence of the splice acceptor site is altered such that it comprises a restriction enzyme site. For example, the altered sequence can be altered such that it comprises an NcoI restriction enzyme site. However, there are a very large number of other restriction enzyme sites that could be provided. A benefit of introduction of a restriction enzyme site at the altered splice acceptor site is that it allows for easy analysis for the occurrence of a successful editing event.
In one embodiment the splice acceptor site is edited to alter the sequence from AATGCTATTTTTCAGCCCACAGGAAACCCAGG (SEQ ID NO: 3) to AATGCTATTTTTCgGCCatggGGAAACCCAGG (SEQ ID NO:4). The sequence changes are shown in lower case.
In preferred embodiments of the present invention the genetically edited swine has improved tolerance or resistance to PRRSV infection. Suitably the animal is resistant to PRRS infection. Deletion of SRCR5 from CD163 has been shown to result in CD163 expressing cells, particularly pulmonary alveolar macrophages (PAMs) and peripheral blood monocyte-derived macrophages (PMMs), becoming highly resistant to infection with PRRSV.
According to a second aspect of the present invention there is provided a genetically edited swine cell or embryo, wherein the edit results in the deletion of SRCR5 domain from CD163 protein produced by the swine cell or embryo. “Cell or embryo” in this context encompasses a somatic cell, germ cell, stem cell, gamete, zygote, blastocyst, embryo, foetus and/or donor cell.
The various features discussed with regard to the first aspect of the invention apply mutatis mutandis to the second aspect of the invention. For example, the nature of the various edits discussed above in respect of the swine apply equally to the edited cell or embryo.
According to a third aspect of the present invention there is provided a method of producing a genetically edited swine, the method comprising the steps of:
The genome modification that results in deletion of SRCR5 from the CD163 protein can be deletion of exon 7 from the CD163 gene or the inactivation of the splice acceptor site associated with exon 7 of the CD163 gene, i.e. the splice acceptor site located at the 5′ end of Exon 7.
In step a) the swine cell can be any suitable cell. Suitably the swine cell can be a somatic cell, a gamete, a germ cell, a gametocyte, a stem cell (e.g. a totipotent stem cell or pluripotent stem cell) or a zygote.
Preferably the method is performed on a zygote. The term ‘zygote’ can be used in a strict sense to refer to the single cell formed by the fusion of gametes. However, it can also be used more broadly in the present context to refer to the cell bundle resulting from the first few divisions of the true zygote—this is more properly known as the morula.
It is preferred that the present method is at least initiated, and preferably completed, in the zygote at the single cell stage. This should result in all cells of the swine containing the same edit. It is, however, possible that the zygote may divide while the editing process is occurring. Depending on when the cell division occurs relative to the stage of the editing process, it is possible that one of the following will occur:
Editing can also be conducted after the first cell division, and the results may be of interest. However, this is generally less preferred where the desired result is a non-mosaic animal.
Step b) suitably comprises:
The editing event that results in deletion of SRCR5 from the CD163 protein can be deletion of exon 7 from the CD163 gene or the inactivation of the splice acceptor site associated with exon 7, i.e. located at the 5′ end of Exon 7.
In certain embodiments step b) suitably comprises introducing site-specific nucleases to the cell which are targeted to target sites flanking exon 7 of the CD163 gene so as to induce double-stranded DNA cuts on either side of exon 7 and thereby cause its deletion. The target sites are suitable in introns 6 and 7. Where a target site is in intron 6, the cutting site is preferably 3′ of the splice donor site at the 3′ end of exon 6. Where a target site is in intron 7, the cutting site is preferably 5′ of the splice acceptor site at the 5′ of exon 8.
In certain embodiments step b) suitably comprises introducing an upstream site-specific nuclease to the cell, the upstream site-specific nuclease targeting a target site upstream of exon 7 of the CD163, and introducing a downstream site-specific nuclease to the cell, the downstream site-specific nuclease targeting a target site downstream of exon 7 of the CD163. ‘Upstream’ in this context refers to a site which is located upstream of the 5′ end of exon 7 of the CD163 gene. Preferably the upstream target site is located in the region between the 5′ end of exon 7 and the splice donor site located at the 3′ end of exon 6. In some embodiments the upstream target site is located within 2000 bases (suitably within 1000 bases, 500 bases, 300 bases, 200 bases or 100 bases) upstream of the 5′ end of exon 7. The cutting site of a site-specific nuclease is typically within or very close to its target site, and thus the site-specific nuclease induces a DNA cut within 2000, 1000, 500, 300, 200 or 100 bases upstream of the 5′ end of exon 7. The cutting site of the site-specific nuclease is suitably in the region between the 5′ end of exon 7 and the splice donor site located at the 3′ end of exon 6.
The skilled person can readily target known site-specific nucleases (such as CRISPR/Cas9 or other CRIPR nucleases, TALENs or ZFNs) to any desired target sited in the regions discussed above. In the case of CRISPR/Cas9 or other CRIPR nucleases the method suitably comprises providing a guide RNA to direct the Cas9 protein to the desired target site. In the case of TALEN or ZFN it is the protein code of the site-specific nuclease that determines the binding site of the site-specific nuclease.
Exemplary upstream target sites which can be used in the case where the site-specific nuclease is CRISPR/Cas9, along with the associated cut location and sgRNAs are given below (cut locations are shown by the “|” symbol):
‘Downstream’ in this context refers to a site which is located at or near the 3′ end of exon 7 of the CD163 gene. Typically, a downstream site is located in intron 7. Preferably the downstream target site is located in the region between the 3′ end of exon 7 and the splice acceptor site located at the 5′ end of exon 8. In some embodiments the downstream target site is located within 75 bases or 50 bases 3′ of the 3′ end of exon 7. The cutting site of the site-specific nuclease is thus suitably within this defined region, so that the cut occurs 3′ of the 3′ end of exon 7, and 3′ of the 5′ end of the splice acceptor site located at the 5′ end of exon 8, for example, the cutting site of the site-specific nuclease is typically 5′ of the splice acceptor site located at the 5′ end of exon 8.
An exemplary downstream target site that can be used in the case where the site-specific nuclease is CRISPR/Cas9, along with the associated cut location and sgRNA sequence are given below (cut location is shown by the “|” symbol):
In certain embodiments step b) suitably comprises introducing a site-specific nuclease that targets the splice acceptor site associated with exon 7, i.e. located at the 5′ end of Exon 7.
Suitably a site-specific nuclease induces a double stranded cut within or near to the splice acceptor site associated with exon 7.
In some embodiments the site-specific nuclease induces a cut in the region extending from position 23378 to position 23416 with reference to SEQ ID NO:1, or at a position within 200, 100, 50 or 25 bases of said region in a 5′ or 3′ direction. In other words, the site-specific nuclease induces a double stranded cut in the predicted splice acceptor site associated with exon 7, or in flanking regions.
The skilled person can readily target known site-specific nucleases (such as CRISPR/Cas9, TALENs or ZFNs) to any desired target site in the regions discussed above. In the case of CRISPR/Cas9 and other CRIPR nucleases, the method suitably comprises providing a guide RNA to direct the Cas9 or other CRIPR nuclease protein to the desired target site. In the case of TALEN or ZFN it is the protein code of the site-specific nuclease that determines the binding site of the site-specific nuclease.
In the case of CRISPR/Cas9 mediated gene editing, suitable guide RNA sequences to target the splice acceptor site associated with exon 7 are as follows:
These two guide sequences result in the induction of double stranded cut sites at the following sequences at the 5′ end of exon 7 by Cas9 (cut locations are shown by the “|” symbol):
The site-specific nuclease suitably creates a single double stranded cut at the desired cutting site. In that case the splice acceptor site associated with exon 7 can be inactivated by non-homologous end joining (NHEJ) or by homology directed repair (HDR). Where HDR is the intended method of inactivation, an HDR template is provided. As is well-known in the art, the HDR template comprises a central portion, which contains the sequence intended to replace the normally occurring sequence, and flanking portions which are homologous to the normal sequence. The HDR template thus suitably comprises a central portion that has a sequence that, when introduced to the CD163 gene by HDR, inactivates the splice acceptor site.
An exemplary, but non-limiting, HDR template has the following sequence: GAAGGAAAATATTGGAATCATATTCTCCCTCACCGAAATGCTATTTTTCgGCCatggGGAAAC CCAGGCTGGTTGGAGGGGACATTCCCTGCTCTGGTC (SEQ ID NO:16) (lower case letters show the changes made compared to the unaltered sequence).
While the exemplary target sites set out above relate to the CRISPR/Cas9 site specific nuclease, it will be immediately apparent to the skilled person that many other target sites could be used, and also that other site specific nucleases (often referred to as ‘editors’ or ‘gene editors’ in this context) could be used. Suitable target sites for alternative site specific nucleases could readily be determined by the skilled person.
In preferred embodiments the site-specific nuclease comprises at least one zinc finger nuclease (ZFN), Transcription Activator-Like Effector Nuclease (TALEN), RNA-guided CRISPR nuclease (e.g. CRISPR/Cas9 or other CRISPR nuclease, such as CRISPR/Cpf), or a meganuclease.
The site-specific nuclease is typically capable of creating a double stranded break in the genomic DNA. This can be achieved with a number of site-specific nucleases, including, but not limited to, CRISPR/Cas or other CRISPR nuclease, ZFNs and TALENs.
In some embodiments the site-specific nuclease comprises a pair of cooperating site-specific nucleases, each of which are able to generate a single stranded break. Suitably the site-specific nuclease comprises a pair of cooperating ZFNs, TALENs or CRISPR ‘nickases’ (e.g. having a modified Cas9 or other nuclease capable of cutting only one DNA strand), which cooperate to generate a double stranded break in the genomic DNA. In such embodiments the target site suitably comprises a pair of half sites, with one of the pair binding at each half site. Thus, in some embodiments the site-specific nuclease comprises a pair of ZFNs, TALENs or RNA-guided CRISPR ‘nickases’ (e.g. having a modified Cas9 or other nuclease capable of cutting only one DNA strand), capable of causing a double stranded DNA break only when both members of the pair are present and form a heterodimer which is able to cut both strands of the DNA molecule. In some preferred embodiments the site-specific nuclease comprises a pair of ZFNs. The use of pairs of corresponding site-specific nucleases can have benefits in reducing off-target cuts
It should be noted that the site-specific nuclease can be introduced to a cell in any suitable form. For example, the nuclease can be provided directly into the cell as a functional protein. Alternatively, the nuclease can be provided into the cell in the form of a precursor or template from which the active nuclease is produced by the cell. In a preferred embodiment an mRNA encoding the nuclease is introduced into the cell, e.g. by injection. The mRNA is then expressed by the cell to form the functioning protein. Using mRNA in this way allows rapid but transient expression of the nuclease within the cell, which is ideal for the purposes of genetic editing. Where an RNA is used to target the site-specific nuclease, this can be provided in any suitable form.
It should also be noted that the term ‘nuclease’ is intended to cover any biological enzyme which creates a single or double stranded cut of a target nucleic acid. Accordingly, the term includes nickases and recombinases, as well as more conventional nucleases which cause single or double stranded breaks.
ZFN technology is described extensively in the literature and, inter alia, in the following patent documents: U.S. Pat. Nos. 6,479,626, 6,534,261, 6,607,882, 6,746,838, 6,794,136, 6,824,978, 6,866,997, 6,933,113, 6,979,539, 7,013,219, 7,030,215, 7,220,719, 7,241,573, 7,241,574, 7,585,849, 7,595,376, 6,903,185, 6,479,626, 8,106,255, 20030232410, and 20090203140, all of which are incorporated by reference. ZFNs can be obtained commercially from Sigma-Aldrich (St. Louis, Mo., US) under the CompoZr® Zinc Finger Nuclease Technology branded products and services.
TALEN technology is described extensively in the literature and, inter alia, in the following patent documents: U.S. Pat. Nos. 8,420,782, 8,470,973, 8,440,431, 8,440,432, 8,450,471, 8,586,363, 8,697,853, EP2510096, U.S. Pat. Nos. 8,586,526, 8,623,618, EP2464750, US2011041195, US2011247089, US2013198878, WO2012/116274, WO2014110552, WO2014070887, WO2014022120, WO2013192316, and WO2010008562, all of which are incorporated by reference. TALENs can be obtained commercially from Thermo Fisher Scientific, Inc. (Waltham, Mass., US) under the GeneArt® TALs branded products and services (formerly marketed under the Life Technologies brand).
CRISPR/Cas technology is described extensively in the literature (e.g. Cong et al. ‘Multiplex Genome Engineering Using CRISPR/Cas Systems’, Science, 15 Feb. 2013: Vol. 339 no. 6121 pp. 819-823) and, inter alia, in the following patent documents: U.S. Pat. No. 8,697,359, US2010076057, WO2013/176772, U.S. Pat. No. 8,771,945, US2010076057, US2014186843, US2014179770, US2014179006, WO2014093712, WO2014093701, WO2014093635, WO2014093694, WO2014093655, WO2014093709, WO2013/188638, WO2013/142578, WO2013/141680, WO2013/188522, U.S. Pat. No. 8,546,553, WO2014/089290, and WO2014/093479, all of which are incorporated by reference. CRISPR/Cas systems can be obtained commercially from Sigma-Aldrich (St. Louis, Mo., US) under the CRISPR/Cas Nuclease RNA-guided Genome Editing suite of products and services, or from Thermo Fisher Scientific, Inc. (Waltham, Mass., US) under the GeneArt® CRISPR branded products and services. CRISPR/Cpf has also been widely described in the literature.
Of course, in this rapidly developing field other techniques for genetic editing are likely to become available. Such techniques could, in many cases, be readily adapted for use in the present invention.
With regard to step c) of, there are a range of well-known techniques in the art that can be used to produce animals from cells comprising genetic alterations. Such techniques include, without limitation, pronuclear microinjection (U.S. Pat. No. 4,873,191) or electroporation of embryos (Lo (1983) Mol. Cell. Biol. 3, 1803-1814), sperm-mediated gene transfer (Lavitrano et al. 25 (2002) Proc. Natl. Acad. Sci. USA 99, 14230-14235; Lavitrano et al. (2006) Reprod. Fert. Develop. 18, 19-23), and in vitro transformation of somatic cells, such as cumulus or mammary cells, or adult, fetal, or embryonic stem cells, followed by nuclear transplantation (Wilmut et al. (1997) Nature 385, 810-813; and Wakayama et al. (1998) Nature 394, 369-374). Standard breeding techniques can be used to create animals that are homozygous or biallelic for a desired genetic edit from initially heterozygous founder animals. The specific description gives details of an exemplary, but not limiting, method for generating animals from an edited zygote. The present invention is not limited to any specific way of generating an animal from the edited cell produced in step b).
Step c) of the method can optionally involve cloning, e.g. somatic cell nuclear transfer (SCNT). In such an embodiment the genetic editing event is carried out on a somatic cell, after which the edited nucleus is transferred to an enucleated egg cell. Typically a population of somatic cells will be edited and cells in which a desired editing event has occurred will be used to provide donor nuclei for SCNT. Processes for SCNT have been well described in the art and would be known to the skilled person. However, it is an advantage of the present invention that editing can be performed without the need for cloning.
The method may suitably comprise crossing a swine produced from the genetically edited cell with another swine to obtain a descendent swine. Preferably the descendent swine is homozygous or biallelic for the genome edit that results in the deletion of SRCR5 from CD163 produced by the animal. This can be achieved, for example, by crossing two heterozygous swine, as is well known in the art. Thus, in some embodiments the method suitably comprises step d), crossing a swine produced in step c) (which can be heterozygous for the genome edit that results in the deletion of SRCR5 from CD163 produced by the animal), with another swine that is heterozygous for the genome edit that results in the deletion of SRCR5 from CD163 produced by the animal.
In certain embodiments, the method of the present invention comprises the steps of:
The genetically edited zygote can be grown to become an embryo and eventually an adult animal. As discussed above, if the editing event occurs in the single-cell zygote then all cells of this animal will therefore comprise the modified CD163 gene as all cells of the animal are derived from a single genetically edited cell. If the editing event occurs after one or more cell divisions then the resultant animal will likely be a mosaic for the editing event, in that it will have some cells derived from the edited cell and some cells derived from unedited cells.
The method may involve characterising the genetic editing event that has occurred. Suitable methods to achieve this are set out below.
The method can be performed on a plurality of zygotes and the method may involve selecting zygotes in which the desired genetic modification has been achieved.
Preferably the swine produced according to the methods of the present invention is homozygous or biallelic for the genome edit that results in the deletion of SRCR5 from CD163 produced by the animal. This can be achieved directly as a result of the editing process of step b), or by a subsequent crossing step between two heterozygous swine.
According to the fourth aspect of the present invention there is provided a method of producing a genetically edited swine cell or embryo, the method comprising the steps of:
The various features discussed with regard to the third aspect of the invention apply to the fourth aspect of the invention mutatis mutandis.
According to a fifth aspect of the invention there is provided an animal, cell or embryo produced according to the third or fourth aspects of the invention.
According to a sixth aspect of the present invention, there is provided a method of modifying a swine to increase its resistance or tolerance to PRRSV comprising editing the genome of cells in the swine to create a modification which results in the deletion of SRCR5 domain of the CD163 protein.
According to a seventh aspect of the present invention, there is provided a swine or a cell of a swine which expresses or bears a CD163 protein in which the SRCR5 domain has been deleted. The cell may suitably be a macrophage, and in some cases can be a peripheral blood monocyte-derived macrophages (PMM) or pulmonary alveolar macrophage (PAM).
Embodiments of the present invention will now be described, by way of non-limiting example, with reference to the accompanying drawings.
While the making and using of various embodiments of the present invention are discussed in detail below, it should be appreciated that the present invention provides many applicable inventive concepts that can be embodied in a wide variety of specific contexts. The specific embodiments discussed herein are merely illustrative of specific ways to make and use the invention and do not delimit the scope of the invention.
To facilitate the understanding of this invention, a number of terms are defined below. Terms defined herein have meanings as commonly understood by a person of ordinary skill in the areas relevant to the present invention. Terms such as “a”, “an” and “the” are not intended to refer to only a singular entity, but include the general class of which a specific example may be used for illustration. The terminology herein is used to describe specific embodiments of the invention, but their usage does not delimit the invention, except as outlined in the claims.
The term “swine”, or variants thereof, as used herein refers to any of the animals in the Suidae family of even-toed ungulates including animals in the genus Sus and other related species, including the peccary, the babirusa, and the warthog.
The term “pig” or variants thereof as used herein refers to any of the animals in the genus Sus. It includes the domestic pig (Sus scrofa domesticus or Sus domesticus) and its ancestor, the common Eurasian wild boar (Sus scrofa). For the present purposes, the domestic pig is considered to be a sub-species of the species Sus scrofa. It does not include the peccary, the babirusa, and the warthog.
The term “domestic pig”, or variants thereof, as used herein refers to an animal of the sub-species Sus scrofa domesticus.
The term “site-specific nuclease”, or variants thereof, as used herein refers to engineered nucleases which can be configured to cut DNA at a desired location. Such site-specific nucleases are also known as engineered nucleases, targetable nucleases, genome editing nucleases, molecular scissors, and suchlike. Examples of site-specific nucleases include zinc finger nucleases (ZFNs), Transcription Activator-Like Effector Nucleases (TALENs), the CRISPR/Cas system (CRISPR/Cas), and meganucleases, such as hybrid meganucleases.
“Genetically edited” or “genetically modified” when used in relation to subject biological material, refers to the fact that the subject biological material has been treated to produce a genetic modification thereof compared to control, e.g. wild type, biological material.
“Target site” refers to the site having a nucleic acid sequence to which a site-specific nuclease binds. When the site-specific nuclease bind at a target site it acts to cut the DNA within or adjacent to the target site (this can be achieved by a single site-specific nuclease, or a corresponding pair or nucleases, in which case there will be two so-called “half-sites”, as desired), the location of the cut being referred to as the “cut site” or “cutting site”. Where a target site is defined for a site-specific nuclease above, the cut site is suitably with the target site, or adjacent to the target site. Where the target site is mentioned as being near or adjacent to a specific feature in the genome, e.g. a feature to be deleted or preserved in an editing event (such as exon 7 or a splice site), the cutting site should be located so as to achieve the desired outcome, i.e. it results in deletion or preservation of the feature, as desired. Site-specific nucleases can be designed to target any desired target site; for example, with CRISPR/Cas9 this can be achieved using a suitable sgRNA, and for ZFN or TALENs suitable proteins can be designed and obtained from commercial sources.
“ΔSRCR5” refers to an animal, typically a swine, which comprises a biallelic or homozygous CD163 SRCR5 deletion.
“Unaltered” with reference to a nucleic acid sequence (such as a region of the genome or a gene) means that the sequence has not been altered from the wild type sequence.
“Tolerance or resistance”—an animal can be said to be more tolerant or resistant to PRRSV infection when the mortality rate, morbidity rate, the proportion of animals showing significant morbidity (e.g. weight loss or decreased growth rate), the level of morbidity or the duration of morbidity is reduced when animals are challenged with PRSSV infection. Any statistically significant reduction (e.g. 95% confidence, or 99% confidence using an appropriate test) in the mortality or morbidity between a population of genetically edited pigs and a population of equivalent non-edited pigs when exposed to PRRSV of the same virulence level (ideally the same isolate) demonstrates improved tolerance or resistance. Improved tolerance or resistance can be demonstrated by a reduced susceptibility to PRRSV inflection, or a lessening of the symptoms when infection occurs. Improved resistance to infection in a swine can be tested in vitro using the methodologies described below for PAM and PMM cells.
“Protein” and “peptide”, as used herein, can be used interchangeably (unless the context suggests otherwise) and mean at least two covalently attached amino acids linked by a peptidyl bond. The term protein encompasses purified natural products, or products which may be produced partially or wholly using recombinant or synthetic techniques. The terms peptide and protein may refer to an aggregate of a protein such as a dimer or other multimer, a fusion protein, a protein variant, or derivative thereof. A protein may comprise amino acids not encoded by a nucleic acid codon, i.e. non-natural amino acids.
PRRS is one of the most economically important infectious diseases affecting pigs worldwide. The “mystery swine disease” was first observed almost simultaneously in North America and in Europe in the late 1980s [1,2]. The causative agent of PRRS was identified to be a virus later named PRRS virus (PRRSV). Infected pigs may present with symptoms involving inappetence, fever, lethargy, and respiratory distress. However, the most devastating effects of PRRSV infection are observed in young piglets and pregnant sows. In pregnant sows an infection with PRRSV can cause a partial displacement of the placenta, leading to full abortions or to death and mummification of fetuses in utero [3]. Late-term abortions occur in up to 30% of infected sows with litters containing up to 100% stillborn piglets. Live-born piglets from an antenatal infection are often weak and display severe respiratory symptoms, with up to 80% of them dying on a weekly basis pre-weaning [4,5]. Young piglets infected with PRRSV often display diarrhea and severe respiratory distress caused by lesions in the lung. In pre-weaned piglets the infection may be transmitted via the mammary gland secretions of an infected sow [6]. At this age the infection has a fatal outcome in up to 80% of animals. After weaning mortality rates reduce, but continued economic losses due to reduced daily gain and feed efficiency are often observed [4,7,8]. Due to reduction or loss of pregnancies, death in young piglets, and decreased growth rates in all PRRSV infected pigs it is estimated that more than $650m are lost annually to pork producers in the United States alone [9,10].
PRRSV is an enveloped, plus-strand RNA virus belonging to the Arteriviridae family in the order Nidovirales [11,12]. The PRRSV genome (˜15 kb) encodes at least 12 non-structural and seven structural proteins. The viral RNA is packaged by the nucleocapsid protein N, which is surrounded by the lipoprotein envelope, containing the non-glycosylated membrane proteins M and E, as well as four glycosylated glycoproteins GP2, GP3, GP4, and GP5, whereby GP2, 3, and 4 form a complex [13-17].
PRRSV has a very narrow host range, infecting only specific subsets of porcine macrophages [18-20]. It is unknown yet how widespread PRRSV infections are within the superfamily of the Suidae. Whereby European wild boars have been shown to act as a reservoir for PRRSV [21], little is known about infection in African suids, such as bushpigs and warthogs. In vitro virus replication is supported by the African Green Monkey cell line MARC-145. Entry of PRRSV into macrophages has been shown to occur via pH-dependent, receptor mediated endocytosis [22,23]. Various attachment factors and receptors have been indicated to be involved in the PRRSV entry process (reviewed in [24]). Heparan sulphate was identified early as an attachment factor of the virus [25-27]. In vitro infection of pulmonary alveolar macrophages (PAMs) but not MARC-145 cells was shown to be inhibited by an antibody targeting CD169 (sialoadhesin), a lectin expressed on the surface of macrophages [28]. Overexpression of CD169 in previously non-permissive PK-15 cells showed internalization but not productive replication of PRRSV [29]. Finally, an in vivo challenge of genetically modified pigs in which the CD169 gene had been knocked out revealed no increased resistance to PRRSV infection, suggesting that CD169 is an attachment factor that is not essential for PRRSV infection [30]. Even though cell surface protein expression is a major determinant of PRRSV binding and internalization, there appears to be a redundancy amongst cell surface attachment factors, with the potential for additional, as yet unidentified receptors, being involved [31]. The scavenger receptor CD163, also known as haptoglobin scavenger receptor or p155, is expressed on specific subtypes of macrophages and has been identified as a fusion receptor for PRRSV. The extracellular portion of CD163 forms a pearl-on-a-string structure of nine scavenger receptor cysteine-rich (SRCR) domains and is anchored by a single transmembrane segment and a short cytoplasmic domain [32]. CD163 has a variety of biological functions, including mediating systemic inflammation and the removal of hemoglobin from blood plasma (reviewed in [33,34]). Overexpression of CD163 renders non-susceptible cells permissive to PRRSV infection [35], whereby it was found that CD163 does not mediate internalization but is crucial for fusion [36]. The transmembrane anchoring and an interaction with the SRCR domain 5 (SRCR5) of CD163 were found to be essential for successful infection with PRRSV [34,35]. Recent in vivo experiments with CD163 knock-out pigs have been performed [37]. However, as CD163 has important biological functions the complete knockout could have a negative physiological impact pigs, particularly with respect to inflammation and/or infection by other pathogens.
This study aimed to generate pigs with a defined CD163 SRCR5 deletion and to assess the susceptibility of macrophages from these pigs to PRRSV infection.
Materials and Methods
All animal work was approved under UK Home Office license after review by the University of Edinburgh's Animal Ethics Committee and was carried out in accordance with the approved guidelines.
Cells and Viruses
Primary pulmonary alveolar macrophages (PAMs) for the propagation of PRRSV genotype 1, subtype 1 strain H2 (PRRSV H2) [52], subtype 2 strain DAI (PRRSV DAI) [53], and subtype 3 strain SU1-Bel (PRRSV SU1-Bel)[54] were harvested from wild type surplus research animals aged 6-9 weeks as previously described [45]. Briefly, animals were euthanized according to a schedule I method. Lungs were removed and transferred on ice to a sterile environment. PAMs were extracted from lungs by washing the lungs twice with warm PBS, massaging to release macrophages. Cells were collected by centrifugation for 10 min at 400 g. When necessary red cells were removed using red cell lysis buffer (10 mM KHCO3, 155 mM NH4Cl, 0.1 mM EDTA, pH 8.0) for 5 min before washing again with PBS. Cells were collected by centrifugation as before and frozen in 90% FBS (HI, GE Healthcare), 10% DMSO (Sigma). Cells were frozen gradually at 1° C./min in a −80° C. freezer before being transferred to −150° C.
PAMs from the animals 627, 628, 629, 630, 633, and 634 were collected at 8 weeks of age. For this the piglets were sedated using a Ketamine/Azaperone pre-medication mix and anaesthetized with Ketamine/Midazolam. Anesthesia throughout the procedure was maintained using Sevoflurane. PAMs were collected by bronchoalveolar lavage (BAL) through an intubation with an air flow access. Three lung segments were flushed in each animal using 2×20 ml PBS. Fluid recovery was between 60-80%. Cells were collected by centrifugation for 10 min at 400 g from the BAL fluid and frozen as above.
Peripheral blood monocytes (PBMCs) were isolated as described previously [45]. Briefly, blood was collected using EDTA coated vacuum tubes from the jugular vein of the piglets at 10 weeks of age. Blood was centrifuged at 1200 g for 15 min and buffy coat transferred to PBS. Lymphoprep (Axis-Shield) was overlaid with an equal volume of buffy coat/PBS and centrifuged for 45 min at 400 g. The mononuclear cell fraction was washed with PBS, cells collected and frozen as described above.
PAM cells were cultivated in RPMI-1640, Glutamax (Invitrogen), 10% FBS (HI, GE Healthcare), 100 IU/ml penicillin and 100 μg/ml streptomycin (Invitrogen) (cRPMI). PBMCs were cultivated in cRPMI supplemented with rhCSF-1 (1×104 units/ml; a gift from Chiron) for 6 days prior to infection.
PK15 cells were cultured in DMEM supplemented with Glutamax (Invitrogen), 10% FBS (HI, GE Healthcare), 100 IU/ml penicillin and 100 μg/ml streptomycin (Invitrogen).
Design and In Vitro Cutting Efficiency Assessment of Guide RNAs
Three potential guide RNA sequences were selected in the 200 bp of intron 6 and one in the 97 bp long intron 7. Oligomers (Invitrogen) were ordered, hybridized as previously described [72] then ligated into the BbsI sites of plasmid pSL66 (a derivative of px458 with modifications to the sgRNA scaffold as described by [42]). The generated plasmids contain a hU6 promoter driving expression of the guide RNA sequence and a CBA promoter driving Cas9-2A-GFP with an SV40 nuclear localization signal (NLS) at the N-terminus and a nucleoplasmin NLS at the C-terminus of Cas9. Cutting efficiency of each guide was assessed by transfection of the plasmids into pig PK15 cells using a Neon transfection system (Invitrogen) set at 1400 mV with 2 pulses of 20 mS. 48 hours post-transfection GFP positive cells were collected using a FACS Aria III cell sorter (Becton Dickinson) and cultured for a further 4 days prior to preparation of genomic DNA (DNeasy Blood & Tissues Kit, Qiagen). PCR across the target sites was with oSL46 (ACCTTGATGATTGCGCTCTT—SEQ ID NO:17) and oSL47 (TGTCCCAGTGAGAGTTGCAG—SEQ ID NO:18) using AccuPrime Taq DNA polymerase HiFi (Life Technologies) to produce a product of 940 bp. A Cell assay (Transgenomic; Surveyor Mutation Detection Kit) was performed as previously described [73]. Co-transfection of PK15 cells with pairs of plasmids encoding guides flanking exon 7 were carried out as described above with the exception that cells were harvested at 40 hours post-transfection without enrichment for GFP expression. In this instance a truncated PCR product was observed in addition to the 940 bp fragment, indicating deletion of exon 7.
Based on both single and double cutting efficiencies guide RNAs SL26 (GAATCGGCTAAGCCCACTGT—SEQ ID NO:7), located 121 bp upstream of exon 7, and SL28 (CCCATGCCATGAAGAGGGTA—SEQ ID NO:11), located 30 bp downstream of exon 7 were selected for in vivo experiments.
Generation of Guide RNA and Quality Assessment
A DNA oligomer fragment containing the entire guide RNA scaffold and a T7 promoter was generated by PCR from the respective plasmid template as follows; a forward primer containing the T7 promoter followed by the first 18 bp of the respective guide RNA and the reverse primers oSL6 (AAAAGCACCGACTCGGTGCC—SEQ ID NO:19) were used in combination with the Phusion polymerase (NEB). DNA fragments were purified on a 1.5% agarose gel using the MinElute Gel Extraction Kit (Qiagen) according to the manufacturer's instructions. DNA eluate was further treated with 200 μg/ml Proteinase K (Qiagen) in 10 mM Tris-HCl pH 8.0, 0.5% SDS for 30 min at 50° C. followed by phenol/chloroform extraction. Guide RNAs were generated from the resultant DNA fragment using the MEGAshortscript Kit (Thermo Fisher) according to the manufacturer's instructions. RNA was purified using phenol/chloroform extraction followed by ethanol precipitation and resuspended in EmbryoMax Injection Buffer (Millipore). Purity and concentration of the RNA was assessed using an RNA Screen Tape (Agilent) on an Agilent TapeStation according to the manufacturer's instructions.
Zygote Injection and Transfers
Embryos were produced from Large White gilts as described previously [73]. Briefly, gilts were superovulated using a regumate/PMSG/Chorulon regime between day 11 and 15 following estrus. Following heat, the donor gilts were inseminated twice in a 6 hour interval. Zygotes were surgically recovered from mated donors into NCSU-23 HEPES base medium, then subjected to a single 2-5 pl cytoplasmic injection with an injection mix containing 50 ng/μl of each guide (SL26 and SL28) and 100 ng/μl Cas9 mRNA (PNA Bio or Tri-Link) in EmbryoMax Injection buffer (Millipore). Recipient females were treated identically to donor gilts but remained unmated. During surgery, the reproductive tract was exposed and 24-39 zygotes were transferred into the oviduct of recipients using a 3.5 French gauge tomcat catheter. Litter sizes ranged from 5-12 piglets.
In Vitro Assessment Genome Editing in Blastocyst
Uninjected control zygotes and injected surplus zygotes are cultivated in NCSU-23 HEPES base medium, supplemented with cysteine and BSA at 38.5° C. for 5-7 days. Blastocysts were harvested at day 7 post cultivation and the genome amplified using the REPLI-g Mini Kit (Qiagen), according to the manufacturer's instructions. Genotyping was performed as described below.
Genotyping
Genomic DNA was extracted from ear biopsy or tail clippings taken from piglets at 2 days postpartum using the DNeasy Blood and Tissue Kit (Qiagen). The region spanning intron 6 to exon 8 was amplified using primers oSL46 (ACCTTGATGATTGCGCTCTT—SEQ ID NO:17) and oSL47 (TGTCCCAGTGAGAGTTGCAG—SEQ ID NO:18), generating a 904 bp product from the intact allele and a 454 bp product if complete deletion of exon 7 had occurred. PCR products were analyzed by separation on a 1% agarose gel and subsequent Sanger sequencing of all truncated fragments. Fragments corresponding to the wild type length were further analyzed by T7 endonuclease I (NEB) digestion according to the manufacturer's instructions.
RNA Phenotyping
RNA was isolated from 1E6 PAM cells, isolated by BAL as described above, using the RNeasy Mini Kit (Qiagen), according to the manufacturer's instructions, including an on-column DNase digestion. First-strand cDNA was synthesized using an Oligo-dT primer in combination with SuperScript II reverse transcriptase (Invitrogen), according to the manufacturer's instructions. The cDNA was used to assess the RNA phenotype across exons 4 to 9 using primers P0083 (ATGGATCTGATTTAGAGATGAGGC—SEQ ID NO:20) and P0084 (CTATGCAGGCAACACCATTTTCT—SEQ ID NO:21), resulting in a PCR product of 1686 bp length for the intact allele and 1371 bp following precise deletion of exon 7. PCR products were analyzed by separation on a 1% agarose gel and subsequent Sanger sequencing of deletion fragments.
Protein Phenotype Analysis by Western Blotting
4E5 PAM cells isolated by BAL were collected by centrifugation at 300 rcf for 10 min. The pellet was resuspended in Laemmli sample buffer containing 100 mM DTT, boiled for 10 min at 95° C. and subjected to electrophoresis on 7.5% acrylamide (Bio-Rad) gels. After transfer to a nitrocellulose membrane (Amersham), the presence of cellular proteins was probed with antibodies against CD163 (rabbit pAb, abcam, ab87099) at 1 μg/ml, and β-actin (HRP-tagged, mouse mAb, Sigma, A3854) at 1:2000. For CD163 the blot was subsequently incubated with HRP-labelled rabbit anti-mouse antibody (DAKO, P0260) at 1:5000. Binding of HRP-labelled antibodies was visualized using the Pierce ECL Western Blotting Substrate (Thermo Fisher), according to the manufacturer's instructions.
Quantification of CD163 mRNA by RT-qPCR
RNA was isolated from 1E6 PAMs using the RNeasy Mini Kit (Qiagen), according to the manufacturer's instructions, including an on-column DNase digestion. RNA levels were measured using the GoTaq 1-Step RT-qPCR system (Promega) according to the manufacturers' instructions on a LightCycler 480 (Roche). mRNA levels of CD163 were quantified using primers P0074 (CATGGACACGAGTCTGCTCT—SEQ ID NO:22) and P0075 (GCTGCCTCCACCTTTAAGTC—SEQ ID NO:23) and reference mRNA levels of β-actin using primers P0081 (CCCTGGAGAAGAGCTACGAG—SEQ ID NO:24) and P0082 (AAGGTAGTTTCGTGGATGCC—SEQ ID NO:25).
Characterization of Macrophages by Flow Cytometry
PAMs were seeded one day prior to analysis. PBMCs were seeded seven days prior to analysis and differentiated by CSF1 stimulation to yield PBMC-derived macrophages (PMMs). Cells were harvested by scraping with a rubber policeman and fixed in 4% formaldehyde/PBS for 15 min at room temperature. Cells were incubated with blocking solution (PBS, 3% BSA) for 45 min before staining with antibodies. Cells were stained with antibodies targeting either mouse anti-pig CD14 (AbD Serotec, MGA1273F, 1:50) and mouse anti-pig CD16 (AbD Serotec, MCA2311PE, 1:200), mouse anti-pig CD169 (AbD Serotec, MCA2316F, 1:50) and mouse anti-pig CD172a (SoutherBiotech, 4525-09, 1:400), mouse anti-human CD151 (AbD Serotec, MCA1856PE, 1:50) and mouse anti-pig SWC9 (CD203a) (AbD Serotec, MCA1973F, 1:50), mouse anti-pig CD163 (AbD Serotec, MCA2311PE, 1:50), or mouse IgG1 or an IgG2b negative control (AbD Serotec, MCA928PE, MCA691F, or Sigma, F6397; same concentration as primary Ab). The cells were washed three times with PBS and resuspended in FACS buffer (2% FBS, 0.05M EDTA, 0.2% NaN3 in PBS). Gene expression determined by antibody labelling was assessed by FACS analysis on a FACS Calibur (Becton Dickinson) using FlowJo software.
High MOI Single-Round Infection Assay
PAMs were seeded one day prior to infection. PBMCs were seeded seven days prior to infection and differentiated by CSF1 stimulation to yield PBMC-derived macrophages PMMs. Cells were inoculated at MOI=1 of the respective virus strain (PRRSV H2, DAI, or SU1-Bel) in cRPMI for 3 h at 37° C. The inoculum was replaced by warm cRPMI. At 19 hpi cells were detached by using a cell scraper. Cells were fixed in 4% Formaldehyde (Sigma-Aldrich) in PBS (Gibco) for 15 min at RT, washed with PBS, and subsequently permeabilized in PBS containing 0.1% Triton-X-100 (Alfa Aesar) for 10 min. Cells were incubated with antibody against PRRSV-N (SDOW17-F, RTI, KSL0607, 1:200) and CD163 (AbD Serotec, MCA2311PE, 1:50) or mouse IgG1 negative controls, as described above, in 3% BSA in PBS. The cells were washed three times with PBS and re-suspended in FACS buffer. Infection levels, determined by antibody labelling, were assessed by FACS analysis on a FACS Calibur (Benson Dickson) using FlowJo software.
Low MOI Multiple-Round Infection Assay
PAMs were seeded one day prior to infection. PBMCs were seeded seven days prior to infection and differentiated by rhCSF1 stimulation to yield PMMs. Cells were inoculated at MOI=0.1 with the respective virus strain (PRRSV H2, DAI, or SU1-Bel) in cRPMI for 3 h at 37° C. Inoculum was removed, cells washed 1× with PBS, and infection continued. At the indicated times post inoculation samples were harvested to be assessed. All samples were frozen and processed once all samples from a time course had been collected.
Viral RNA (vRNA) was extracted from the supernatant samples using the QIAmp Viral RNA Mini Kit according to the manufacturer's instructions. The viral RNA levels were quantified by RT-qPCR using the GoTaq Probe 1-Step RT-qPCR system (Promega) for PRRSV H2 and SU1-Bel and the GoTaq 1-Step RT-qPCR system (Promega) for PRRSV DAI, according to the manufacturer's instructions. For this the following primers and probes were used: H2 fwd (GATGACRTCCGGCAYC—SEQ ID NO:26), H2 rev (CAGTTCCTGCGCCTTGAT—SEQ ID NO:27), H2 probe (6-FAM-TGCAATCGATCCAGACGGCTT-TAMRA—SEQ ID NO:28), (optimal H2 primer/probe sequences obtained from JP Frossard, AHVLA), SU1-Bel fwd (TCTTTGTTTGCAATCGATCC—SEQ ID NO:29), SU1-Bel rev (GGCGCACTGTATGACTGACT—SEQ ID NO:30), SU1-Bel probe (6-FAM-CCGGAACTGCGCTTTCA-TAMRA—SEQ ID NO:31), DAI fwd (GGATACTATCACGGGCGGTA—SEQ ID NO:32), DAI rev (GGCACGCCATACAATTCTTA—SEQ ID NO:33). RNA levels were measured on a LightCycler 480 (Roche) using a standard curve generated from vRNA isolates of high titer stocks.
Infectivity of the virus produced was assessed using a TCID50 assay of selected time points on PAMs isolated from wild type surplus research animals.
mRNA and Protein Levels of Heme Oxygenase 1 Upon Hb-Hp Stimulation of PMMs
PBMCs were seeded seven days prior to analysis and differentiated by CSF1 stimulation to yield PMMs. Hemoglobin (Hb, Sigma-Aldrich, AO, H0267) and Haptoglobin (Hp, Sigma Aldrich, Phenotype 2-2, H9762) were mixed in a 1:1 wt/wt ratio in PBS for 15 min on a vertical roller before experimentation. PMMs were incubated with 100 μg/ml Hb-Hp in cRPMI for 24 h at 37° C. Cells were harvested by scraping with a rubber policeman. RNA was isolated from 1E6 cells using the RNeasy Mini Kit (Qiagen), according to the manufacturer's instructions, including an on-column DNase digestion. RNA levels were measured using the GoTaq 1-Step RT-qPCR system (Promega) according to the manufacturers' instructions on a LightCycler 480 (Roche). mRNA levels of heme oxygenase 1 (HO-1) were quantified using primers P0239 (TACATGGGTGACCTGTCTGG—SEQ ID NO:34) and P0240 (ACAGCTGCTTGAACTTGGTG—SEQ ID NO:35) and reference mRNA levels of β-actin using primers P0081 and P0082. For analysis of protein levels of HO-1 cells were collected by centrifugation at 300 rcf for 10 min. The pellet was re-suspended in Laemmli sample buffer containing 100 mM DTT, boiled for 10 min at 95° C. and subjected to electrophoresis on 12% acrylamide (Bio-Rad) gels. After transfer to a nitrocellulose membrane (Amersham), the presence of cellular proteins was probed with antibodies against HO-1 (mouse mAb, abcam, ab13248, 1:250), and calmodulin (rabbit mAb, abcam, ab45689, 1:1000). The blot was subsequently incubated with HRP-labelled goat anti-rabbit antibody (DAKO, PI-1000) at 1:5000. Binding of HRP-labelled antibodies was visualized using the Pierce ECL Western Blotting Substrate (Thermo Fisher), according to the manufacturer's instructions.
Quantification and Visualization of Hemoglobin-Haptoglobin Uptake
PBMCs were seeded seven days prior to analysis and differentiated by CSF1 stimulation to yield PMMs. For fluorescence microscopy, cells were seeded on glass cover slips. Hemoglobin (Sigma-Aldrich, AO, H0267) was labeled with Alexa Fluor 488 (AF-488) using a protein labelling kit (Molecular Probes) according to the manufacturer's instructions. HbAF488 and Hp were mixed in a 1:1 wt/wt ratio in PBS for 15 min on a vertical roller before experimentation. PMMs were incubated with 10 μg/ml HbAF488-Hp in cRPMI for 30 min at 37° C.
For quantification by FACS the cells were collected with a rubber policeman and washed three times with Ca2+/Mg2+-free PBS to remove surface bound HbAF488-Hp as described previously [60]. Cells were fixed in 4% (wt/v) formaldehyde (Sigma-Aldrich) in PBS (Gibco) for 15 min at RT, washed with PBS, and subsequently permeabilized in PBS containing 0.1% Triton-X-100 (Alfa Aesar) for 10 min. Cells were stained with mouse anti pig CD163 antibody (AbD Serotec, MCA2311PE, 1:50) as described above then washed three times with PBS and re-suspended in FACS buffer. Gene expression determined by antibody labelling was assessed by analysis on a FACS Calibur (Becton Dickinson) using FlowJo software.
For immunofluorescence imaging cells were washed three times with Ca2+/Me-free PBS and fixed in 4% formaldehyde (Sigma-Aldrich) in PBS (Gibco) for 15 min at RT, washed with PBS, then permeabilized in PBS containing 0.1% Triton-X-100 (Alfa Aesar) for 10 min. Cells were washed with PBS and incubated with antibody against CD163 (rabbit pAb, abcam, ab87099, 5 μg/ml) in blocking buffer (PBS, 3% FBS) for 1 h, washed, and incubated with secondary goat anti-rabbit AF594 antibody (A11037, 1:100), AF647 phalloidin (A22287, 1:100), and DAPI (1:10,000; all Life Technologies). The samples were analyzed using a confocal laser-scanning microscope (Zeiss LSM-710).
Immunofluorescence Analysis of RTC Formation in Infected PAMs
PAMs were seeded onto coverslips one day prior to infection. Cells were inoculated at MOI=2 of the respective virus strain (PRRSV H2, DAI, or SU1-Bel) in cRPMI for 3 h at 37° C. The inoculum was replaced by warm cRPMI. At 19 hpi cells were fixed in 4% formaldehyde (Sigma-Aldrich) in PBS (Gibco) for 15 min at RT, washed with PBS, and permeabilized as described above. Cells were washed with PBS and incubated with antibody against PRRSV nsp2 (A gift from Ying Fang, South Dakota State University, [74], 1:400) in blocking buffer for 1 h, washed, and incubated with secondary goat anti-mouse AF488 antibody (A11029, 1:100), AF568 phalloidin (A12380, 1:100), and DAPI (1:10,000; all Life Technologies). The samples were analyzed using a confocal laser-scanning microscope (Zeiss LSM-710).
Results
Generation of Live CD163 SRCR5 Deletion Pigs by CRISPR/Cas9 Editing in Zygotes
The CD163 gene is not correctly represented in the current pig reference genome sequence (Sscrofa10.2) [38]. Through targeted sequencing we have established a detailed model of the porcine CD163 locus (unpublished results L. Zen/A. Archibald/T. Ait-Ali)—the genomic sequence of the CD163 gene is set out below as SEQ ID NO:1. Briefly, CD163 is encoded by 16 exons with exons 2-13 predicted to encode the SRCR domains of the protein [39]. Interestingly, SRCR5 is predicted to be encoded by one single exon, namely exon 7 (
All four sequences were assessed in vitro for cutting efficiency by transfection of porcine kidney PK15 cells with a plasmid based on px458 [42] encoding the complete single guide sequence (sgRNA), driven by the hU6 promoter, and a CAG promoter driving NLS-Cas9-2A-GFP. Transfected cells were isolated by fluorescence activated cell sorting (FACS) for GFP and cutting efficiency at the target site was assessed using a Cell surveyor assay. Three out of four guides were shown to direct cutting of DNA as anticipated (2 upstream and one downstream of exon 7). Following double transfection assay and subsequent PCR analysis it was found that the combination of guides SL26 and 51_28 effectively generated the exon 7 deletion in the CD163 gene (
sgRNAs SL26 and SL28 were microinjected together with mRNA encoding the Cas9 nuclease into the cytosol of zygotes. Editing efficiency was assessed in a small number of injected zygotes by in vitro culture to the blastocyst stage, genomic DNA extraction, whole genome amplification and PCR amplification across exon 7. The analysis revealed that two out of 17 blastocysts contained a deletion of the intended size and Sanger sequencing confirmed the deletion of exon 7. Edited blastocyst B2 showed a clean deletion and subsequent re-ligation at the cutting sites of sgSL26 and sgSL28, whilst edited blastocyst B14 showed that in addition to the intended deletion there was also a random insertion of 25 nucleotides at the target site. None of the full length PCR products showed nucleotide mismatches at either cutting site in a T7 endonuclease assay. The editing rate in the blastocysts corresponds to an overall editing rate of 11.7%.
To generate live pigs, 24-39 zygotes injected with sgSL26, sgSL28, and Cas9 mRNA were transferred into the oviduct of recipient gilts. A total of 32 live piglets were born and genotyping of ear and tail biopsies revealed that four of the piglets had an exon 7 deletion, corresponding to 12.5% of the total. In addition to the intended deletion of exon 7, three out of the four animals showed insertion of new DNA at the target site probably as a consequence of non-homologous end joining repair. Pig 347 showed a 2 bp truncation at the sgSL26 cutting site and a 66 bp insertion between the cutting sites, pig 346 showed a deletion of 304 bp after the cutting site of sgSL26, and pig 310 showed a short 9 bp insertion (having the sequence TCAGTCACT) at the cutting sites. Pig 345 was found to have a precise deletion of exon 7 without insertion or deletion of random nucleotides at the cut sites (
Genotype and Phenotype of F1 Generation Pigs
To generate fully homozygous and heterozygous pigs, 310 was mated with 345. This mating yielded a litter of 6 heterozygous, 2 biallelic/homozygous CD163 SRCR5 deletion (ΔSRCR5), and 4 wild type CD163 piglets (
Animals 627, 628, 629, 630, 633, and 634 were selected for further analysis, representing the various genotypes (wild type, heterozygous, and biallelic/homozygous) and genders. Growth rates of both ΔSRCR5 and heterozygous animals were comparable to wild type animals (Table 1). Blood samples were taken from all six animals at 10 weeks of age and analyzed by a full blood count conducted by the diagnostics laboratory at the Royal (Dick) School of Veterinary Studies, University of Edinburgh. The blood counts of all animals were within reference values (Table 1). Size, stature and other morphological features of ΔSRCR5 and heterozygous pigs were comparable to their wild type siblings (
At 8 weeks of age, pulmonary alveolar macrophages (PAMs) were isolated from all six animals by bronchoalveolar lavage (BAL). DNA was extracted from the PAMs and analyzed by PCR and Sanger sequencing. The PAM genotype confirmed the results obtained from the ear biopsies; 628 and 633 were wild type, 627 and 633 heterozygous, and 629 and 630 ΔSRCR5, respectively. Sequencing of PCR products confirmed that all editing events had resulted in complete deletion of exon 7. Whilst pigs 627 and 633 had a clean deletion of exon 7 with precise re-ligation at the sgSL26 and sgSL28 cutting sites in one allele, 629 had one allele with a clean deletion and one allele with a 9 bp insertion between the sites, and pig 630 had both alleles with the 9 bp insertion. RNA was extracted from the PAMs, converted into cDNA using oligo(dT) primed reverse transcription, amplified by PCR and analyzed by Sanger sequencing. PCR products spanning exons 4 to 9 showed the expected 315 bp deletion in both heterozygous and ΔSRCR5 animals (
Pulmonary Alveolar Macrophages of ΔSRCR5 Pigs are Fully Differentiated and Express Macrophage-Specific Surface Proteins
PAMs isolated by BAL were characterized for the expression of macrophage-specific surface proteins. CD14 and CD16 are not expressed on monocytes but levels increase upon maturation into macrophages. In PAMs CD14 is found at moderate levels, whilst CD16 is strongly expressed [44]. CD14/CD16 staining of the PAMs from the ΔSRCR5, heterozygous, and wild type animals were all within the previously observed and documented levels [45], with difference being observed between the various genotypes (data not shown). CD172a, or also known as SIRPα, is expressed at high levels on both monocytes and macrophages [46] and was expressed at high levels in cells from all animals. CD169, described as an attachment factor for PRRSV [29], is not expressed in monocytes but is highly expressed in tissue macrophages [47] and was expressed at expected levels in cells from our animals (data not shown). As in humans, expression of CD163 in pigs is restricted to monocytes and macrophages. CD163 is expressed at high levels in tissue macrophages, but at low levels in blood monocytes and in bone marrow-derived macrophages [48] (porcine macrophage markers are reviewed in [49]). Both the wild type and the SRCR5 deletion CD163 were recognized on the surface of the PAMs (
ΔSRCR5 Pulmonary Alveolar Macrophages are not Susceptible to Infection with PRRSV Genotype 1
PRRSV has two different genotypes with distinct geographic distribution, with genotype 1 being found primarily in Europe and Asia and genotype 2 in the Americas and Asia. The two genotypes show differences in both antigenicity and severity of pathology and have >15% genome divergence between them (reviewed in [50]). Genotype 1 can be further divided into three subtypes, based on the ORF7 sequence and geographical distribution, whereby subtype 1 is pan-European whilst subtypes 2 and 3 are currently limited to Eastern Europe [51]. Here we tested all genotype 1 subtypes of PRRSV, represented by subtype 1 strain H2 (PRRSV H2) [52], subtype 2 strain DAI (PRRSV DAI) [53], and subtype 3 strain SU1-Bel (PRRSV SU1-Bel) [54], originally isolated from the UK, Lithuania, and Belarus, respectively.
PAMs were infected at an MOI=1 in a single-round infection. 19 hours post inoculation (hpi) the cells were harvested and stained with a FITC-labelled antibody against PRRSV-N protein. Infection levels were assessed by FACS analysis. All three virus subtypes resulted in infection levels of 40-60% in wild type and heterozygous animals, with more than 98% of infected cells being classified as CD163 positive. A slightly higher, statistically significant infection was observed in heterozygous animals infected with PRRSV H2 and DAI. The reason for this is unclear, but may reflect either altered CD163 protein expression profile in heterozygous animals or other, as yet unidentified, genetic properties. By contrast, cells from both ΔSRCR5 animals (629 and 630) were found to be highly resistant to infection in this assay (
Peripheral Blood Monocytes from ΔSRCR5 Pigs Differentiate into CD163-Expressing Macrophages Upon CSF1-Induction and Express Macrophage-Specific Markers
To assess the differentiation potential of monocytes into CD163-expressing macrophages we isolated peripheral blood monocytes (PBMCs) from whole blood then were differentiated them into macrophages by CSF1-induction for seven days. Expression of macrophage specific markers was assessed by immunofluorescence labelling and FACS analysis. CD14 and CD16 levels are clear indicators of the differentiation of peripheral blood monocytes with levels of both increasing significantly upon differentiation [44,46]. In addition to CD172a, CD169, and CD163, whose roles as macrophage markers are discussed above, we included a PBMC differentiation marker, SWC9, also known as CD203a, and the putative PRRSV attachment factor CD151 [55,56].
CD14/CD16 staining of the PMBC-derived macrophages (PMMs) from the ΔSRCR5, heterozygous, and wild type animals were all within the previously observed and documented levels, with no difference being observed between the genotypes (
ΔSRCR5 Peripheral Blood Monocyte-Derived Macrophages Still Function as CD163-Dependent Hemoglobin-Haptoglobin Scavengers.
In addition to its contribution to PRRSV susceptibility, CD163 has been described to have a variety of important biological functions. CD163 is an erythroblast binding factor, enhancing the survival, proliferation and differentiation of immature erythroblasts, through association with SRCR domain 2 and CD163 expressing macrophages also clear senescent and malformed erythroblasts. SRCR domain 3 plays a crucial role as a haemoglobin (Hb)-haptoglobin (Hp) scavenger receptor. Free Hb is oxidative and toxic; once complexed with Hp is cleared through binding to SRCR3 on the surface of macrophages and subsequent endocytosis. This prevents oxidative damage, maintains homeostasis, and aids the recycling of iron. CD163 expressing macrophages were also found to be involved in the clearance of a cytokine named TNF-like weak inducer of apoptosis (TWEAK), with all SRCRs apart from SRCR5 being involved in this process [57]. Soluble CD163 can be found at a high concentration in blood plasma but its function in this niche is still unknown (reviewed in [34,58]). Maintaining these biological functions is likely to be important to the production of healthy, genetically edited animals. Interestingly, none of the biological functions assigned to CD163 have yet been linked to SRCR5. In order to confirm whether ΔSRCR5 macrophages were still able to take up Hb-Hp complexes we performed a variety of in vitro experiments. Hb-Hp complex uptake in PMMs in vitro has been investigated extensively in the past, with PMMs able to take up both Hb and Hb-Hp complexes in a CD163-dependent manner and the inducible form of heme oxygenase, heme oxygenase 1 (HO-1), being upregulated upon Hb-Hp uptake [59,60].
PBMCs were differentiated into PMMs by CSF1-induction for seven days, following which PMMs were incubated in the presence of Hb-Hp for 24 h to stimulate HO-1 upregulation. The HO-1 mRNA upregulation, assessed by RT-qPCR, increased 2- to 6-fold in the PMMs from all animals (
Peripheral Blood Monocyte-Derived Macrophages from ΔSRCR5 Animals are not Susceptible to Infection with PRRSV Genotype 1
To explore the possibility that PMMs could be a suitable alternative to monitor PRRSV infection and investigate whether ΔSRCR5 PMMs, like PAMs, are resistant to PRRSV infection we tested infectivity with all three genotype 1 subtypes of PRRSV, represented by the strains described above.
PMMs were infected at an MOI=1 in a single-round infection. 19 hpi cells were harvested and stained with a FITC-labelled antibody against PRRSV-N protein, with infection levels assessed by FACS. All three subtypes showed infection levels of 35-80% in wild type and heterozygous animals. As observed in PAMs, a slightly higher, statistically significant infection was observed in heterozygous animals infected with PRRSV H2, whilst no significant infection was observed in the cells from ΔSRCR5 animals (
The Arrest in Infection of ΔSRCR5 Pulmonary Alveolar Macrophages (PAMs) Occurs Prior to the Formation of the Replication/Transcription Complex.
In the porcine kidney cell line PK-15, lacking CD163 expression, transfected with the PRRSV attachment factor CD169 the virus was found to be internalized but not to undergo uncoating [36]. This indicates that CD163, in a close interplay with attachment/internalization factors, plays a major role in the entry process of PRRSV. To assess whether the infection process in ΔSRCR5 macrophages is arrested prior to replication we inoculated PAM cells with all three PRRSV genotype 1 subtypes, represented by the strains described above, at MOI=2. The inoculum was removed 3 hpi and infection allowed to continue up to 22 hpi. Cells were fixed and stained for the replication-transcription complexes (RTC) formed by PRRSV upon replication initiation. PRRSV nsp2 protein, involved in the formation of double membrane vesicles (reviewed in [61]) was chosen as a representative marker for the RTC. The cells were permeabilized and stained for the presence of PRRSV nsp2. We found that macrophages from both the wild type and the heterozygous animals infected with PRRSV formed RTCs, independent of the subtype. However, in the macrophages from ΔSRCR5 animals no RTC formation was observed. This underlines the involvement of CD163 in the entry and uncoating process of PRRSV infection. It also supports the deletion of SRCR5 as an effective method to abrogate PRRSV infection before the virus or viral proteins are amplified (
The results of this study show that live pigs carrying a CD163 SRCR5 deletion are healthy and maintain the main biological functions of the protein, whilst the deletion renders target cells of PRRSV resistant to infection with the virus. By using two sgRNAs flanking exon 7 of CD163 in CRISPR/Cas9 editing in zygotes we achieved excision of said exon from the genome of pigs yielding a CD163 ΔSRCR5 genotype. The expression of the truncated gene was confirmed by PCR of cDNA, RT-qPCR and western blotting against CD163. Macrophages isolated from the lungs of wild type CD163, heterozygous and ΔSRCR5 animals showed full differentiation and expression of macrophage surface markers characteristic of macrophages isolated from the pulmonary alveolar areas. PAMs are the primary target cells of PRRSV infection. Assessing infection of PAMs from the different genotype animals in both high dose, single-round infections and low dose, multiple-round infections showed PAMs from ΔSRCR5 pigs to be resistant to infection in vitro. The differentiation ability of cells of the monocytes/macrophages lineage from genetically edited CD163 animals was further confirmed by isolation and differentiation of PBMCs. PMMs from ΔSRCR5 pigs were also shown to be resistant to PRRSV infection. PMMs have a crucial biological role, serving as scavengers for Hb-Hp complexes in the blood. Using uptake experiments of fluorescently labelled Hb-Hp complexes as well as gene upregulation assays to monitor the increase of HO-1 upon Hb-Hp stimulation we confirmed that this important biological function is maintained in macrophages isolated from ΔSRCR5 animals.
Using CRISPR/Cas9 editing in zygotes generated live pigs with exon 7 CD163 deletions. Editing efficiency was highly variable, dependent on surgery days, in both in vitro cultivated blastocysts as well as born animals, whereby it needs to be considered that overall numbers are low. The reagents used on the various surgery days were the same and both insemination and surgery times were kept consistent. However, there are many elements in the genome editing process that rely on highly skilled personnel and technical reproducibility. Recent developments in nucleic acid delivery methods for genome editing in zygotes may offer possible solutions to standardize the genome editing process. Various groups recently reported successful genome editing by in vitro electroporation of CRISPR/Cas9 regents into zygotes isolated from mice and rats without removing the zona pellucida [62-64]. Using electroporation to deliver genome editing reagents in vivo Takahasi et al. showed high success with this method in mouse embryos after 1.6 days of gestation [65]. Use of in vitro electroporation could standardize the injection process and reduce the requirement for highly trained personnel. As an alternative, in vivo electroporation would remove both the requirement for donor animals and the long handling process of zygotes prior to re-implantation, however this procedure has currently only been developed for mice (reviewed in [66]). Three out of four of the founder animals were found to be edited in a mosaic pattern. In animal 310 the mosaicism seems to result from a delayed activity of the CRISPR/Cas9 complex, resulting in an edit of one allele in a single cells at the 4- or 8-cell stage. In animals 345 and 347 an initial editing event appears to occur in one allele at the 1-cell stage and a second editing event, modifying the second allele in one of the cells at the 2-cell stage, resulting in homozygous/heterozygous mosaic animals. Mosaicism has been observed in various studies employing injection of genome editors into porcine zygotes [67-69]. Asymmetric spreading of introduced mRNA seems unlikely following results of Sato et al., who performed in vitro EGFP mRNA injections using parthenogenetically activated porcine oocytes, whereby a relatively homogenous fluorescence pattern could be observed [69]. Rather, mosaicism seems to result from Cas9 protein/sgRNA complexes remaining active throughout several cell divisions or delayed mRNA expression possibly triggered by cell division. The former theory is supported by the genotype of 345 and 347, which very likely have developed from an initial editing step in one allele at the one cell stage and editing of the second allele in one of the 2-cell or 4-cell stage cells. To generate more biallelic animals by direct injection of zygotes, a more active reagent set may be beneficial. Recent studies indicate that injection of Cas9/sgRNA ribonucleoproteins (RNPs) is more efficient than mRNA injection. Also, RNP injection can be combined with in vitro electroporation [70].
The mating of the F0 generation animals 310 and 345 resulted in wild type, heterozygous and biallelic edited animals. This showed that despite mosaicism both animals are germline heterozygous. None of the offspring showed any adverse effect from the genome editing under standard husbandry conditions. Interestingly, one of the animals, 630, displayed a putative gene conversion event. Based on the mechanism of interallelic gene conversion we assume that a homologous recombination occurred in this animal between one allele showing the edited genotype of 345 and the other allele the edited genotype of 310. The gene conversion appears to have occurred at the zygote stage, rendering 630 homozygous for the genotype of 310 (reviewed in [71]).
PRRSV shows a very narrow host cell tropism, only infecting specific porcine macrophage subsets. Isolating these cells from our genetically edited animals and their wild type siblings we showed that removal of the CD163 SRCR5 domain results in complete resistance of the macrophages towards PRRSV infection. We further demonstrated that ΔSRCR5 animals are resistant to infection with all European subtypes of genotype 1. This shows that a targeted removal of SRCR5 is sufficient to achieve complete resistance to PRRSV infection in vitro. PRRSV attachment factors CD151 and CD169 are still expressed on ΔSRCR5 macrophages underlining that these proteins are not sufficient for PRRSV infection. PRRSV infection on macrophages from the ΔSRCR5 animals was halted before the formation of replication transcription complexes proving CD163 to be involved in the entry or uncoating stage of the PRRSV replication cycle. The ΔSRCR5 macrophages will provide a new tool to study this process in detail in a true-to-life system.
As there could be a genetic variation of CD163 within the Suidae superfamily we performed an in vitro control experiment to assess the susceptibility of warthog (Phacocherus africanus) PMMs to PRRSV infection. Interestingly, warthog PMMs were found to be as susceptible to infection with all PRRSV genotype 1 subtypes as the pig PMMs. They all replicated the virus at a similar rate and to comparable titers (data not shown). This indicates that genetic variation of CD163 within the Suidae superfamily is probably very limited and PRRSV infection may be widespread. This also shows that the virus poses a threat to African pig breeding countries. The ΔSRCR5 animals have several advantages over previously described genome edited animals resistant to PRRSV infection. Whitworth et al. generated animals with a premature stop codon in exon 3 of the CD163 gene, resulting in an ablation of CD163 expression [37]. In contrast to this we have demonstrated that specific application of genome editing tools in vivo can be used to efficiently generate animals with precise deletion of exon 7 of CD163, and that these animals retain expression of the remainder of the CD163 protein on the surface of specific differentiated macrophages in a native conformation. We further showed that the macrophages from these ΔSRCR5 animals retain full differentiation potential, both in PAMs as well as PBMCs stimulated to differentiate by CSF-1 addition, and that macrophages from edited animals retain the ability to perform crucial biological functions associated with CD163 expression, such as hemoglobin/haptoglobin uptake. Overall, this study demonstrates that it is possible to utilize a targeted genome editing approach to render swine resistant to PRRSV infection, whilst retaining biological function of the targeted gene. Introduction of CD163 SRCR5 deletion animals in pig breeding could significantly reduce the economic losses associated with PRRSV infection.
Inactivation of Splice Acceptor Site in Intron 6
An alternative strategy to delete the SRCR5 domain of CD163 is to inactivate the splice acceptor site located at the 5′ end of exon 7 in the CD163 gene.
Inactivation of the splice acceptor site in exon 7 can be achieved in a number of ways, and two suitable strategies are discussed briefly below, one involving creating a double stranded cut followed by non-homologous end joining (NHEJ), and the other using homology directed repair (HDR). The first option suitably uses a single guide RNA and NHEJ by the target cell. Using the second approach, HDR, a template is provided which is used by the cell's double strand break repair machinery to introduce a sequence modification. Thereby some nucleotides will be replaced in order to destroy the splice acceptor site in a targeted manner, whilst introducing a restriction site (in the example NcoI) which allows for convenient confirmation that the HDR event has taken place.
Suitable methodologies for achieving editing events in pig embryos and generation of animals from edited embryos are discussed above, and are also extensively discussed in the literature, and thus for conciseness they will not be repeated here.
In the case of CRISPR/Cas9 mediated gene editing, suitable guide RNA sequences to target the splice acceptor site are as follows:
These two guide sequences result in the induction of double stranded cut sites at the following sequences at the 5′ end of exon 7 by Cas9:
Approach 1—NHEJ
An RNP complex of sgRNA1 or 2 with Cas9 binds to the target site in the CD163 gene and causes a double-strand break. Where the break occurs NHEJ events arise, commonly resulting in and insertion of deletion event. It is highly likely that either insertion or deletion events will result in the inactivation of the intron 6 splice acceptor site. It is thereafter simply a matter of identifying embryos having the requisite disabling of the splice acceptor site.
Approach 2—HDR
Again, an RNP complex of sgRNA1 or 2 with Cas9 binds to the target site in the CD163 gene and causes a double-strand break. In this case, however, an HDR template is provided, for example a single or double stranded DNA molecule, which comprises a sequence which results in a change of the sequence in the CD163 gene from:
A suitable HDR template has the following sequence: GAAGGAAAATATTGGAATCATATTCTCCCTCACCGAAATGCTATTTTTCgGCCatggGGAAAC CCAGGCTGGTTGGAGGGGACATTCCCTGCTCTGGTC (SEQ ID NO:16—lower case letters show the changes made compared to the unaltered sequence).
The converted sequence in the context of CD163 results in inactivation of the splice acceptor site and the introduction of the NcoI restriction site. The presence of the NcoI site facilitates identification of embryos/animals in which the desired HDR edit has been achieved.
Further Experimental Work
Genome Editing in Zygotes for ΔSRCR5 CD163 Pigs and Breeding for a Genotypically Uniform F2 Generation
Founder generation F0 animals carrying a deletion of exon 7 in the CD163 gene, which encodes the scavenger receptor cysteine-rich domain 5 (SRCR5) of the protein, were generated by CRISPR/Cas9 gene editing as described above (see also 75). Therefore, zygotes were microinjected with two guide RNAs, sgSL26 and sgSL28, in combination with Cas9 mRNA to achieve CRISPR/Cas9-mediated double-strand breaks (DSBs) flanking exon 7. Subsequent DSB repair lead to a deletion of exon 7 (
As previously described, ΔSRCR5 animals express the ΔSRCR5 CD163 mRNA and protein at equivalent levels to wildtype siblings. Furthermore, native-structure ΔSRCR5 CD163 is recognized on the surface of pulmonary alveolar macrophages (PAMs) by a respective antibody. We have further analyzed whether template-based protein structure prediction using RaptorX confirms these findings towards proper folding of the subdomains and the complete ΔSRCR5 CD163 protein (39). As seen in
Previously, we have shown that PAMs and in vitro differentiated peripheral blood monocytes are resistant to infection with both, porcine reproductive and respiratory syndrome virus 1 (PRRSV-1) and PRRSV-2. Now, we aimed to confirm the in vitro results by assessing resistance towards PRRSV-1 infection in vivo. Therefore, we selected four homozygous ΔSRCR5 F2 animals and four wildtype siblings and semi-siblings. The animals were co-housed from weaning. At 6 weeks of age they were transferred to the specific pathogen-free (SPF) unit and co-housed for the duration of the challenge (
ΔSRCR5 Pigs Show Normal Whole Blood Counts and Soluble CD163 Serum Levels
Prior to being moved to the SPF unit blood samples were taken from all eight pigs and analyzed by a full blood count conducted by the diagnostics laboratory at the Royal (Dick) School of Veterinary Studies, University of Edinburgh. The blood counts of all animals were within reference values indicating good general health and the absence of infection or inflammation. Furthermore, the hemoglobin levels of all animals were within reference values, indicating normal function of the hemoglobin/haptoglobin scavenging activity of CD163 (Table 2).
Serum was collected from all animals prior to movement to the SPF unit and on day 0 prior to challenge with PRRSV-1. The soluble CD163 (sCD163) serum levels were assessed using a commercially available enzyme-linked immunosorbent assay (ELISA) recognizing soluble porcine CD163. Serum CD163 levels were found to be 463.5±68.99 ng/ml in ΔSRCR5 pigs and 433.2±69.57 ng/ml in wildtype pigs (
ΔSRCR5 Pigs Show No Signs of PRRSV-1 Infection
At 7-8 weeks of age the pigs were inoculated intranasally with the PRRSV-1, subtype 2 strain BOR-57 (77). Generally, infections with PRRSV-1, subtype2 strains are associated with mild respiratory symptoms, elevated body temperature, extensive lung pathology and high viremia. The challenge was conducted for a period of 14 days following inoculation at day 0 and day 1 with 5E6 TCID50 of the virus each. Rectal temperature, respiratory and other potential symptoms, and demeanor were recorded each day and serum samples were collected on day 0 (prior to challenge), 3, 7, 10, and 14 (prior to euthanasia). Weights were recorded on day 0, 7, and 14 (prior to euthanasia). People conducting the challenge and analyzing the pathology were blind to the genotype of the animals.
The rectal temperature showed significant elevations on days 6-9 of the challenge in the wildtype animals, whereas no body temperature increase was observed in the ΔSRCR5 animals (
Overall, no signs of infection were detected in ΔSRCR5 animals despite the high inoculation volume and exposure to infected and shedding wildtype animals showing that ΔSRCR5 animals are resistant to PRRSV-1 infection, confirming the results found in vitro with both PRRSV-1 and PRRSV-2.
ΔSRCR5 Pigs Show No Cytokine Response to PRRSV-1 Infection and Generally Normal Cytokine Levels
To assess the inflammation and infection response following PRRSV-1 infection a panel of 20 cytokines were analyzed towards their level in the serum of the pigs. Therefore, we used commercial quantitative antibody arrays and serum samples collected on day 0 (prior to challenge), 3, 7, 10, and 14 of the challenge. Overall, cytokine levels on day 0, considered a baseline, were similar between ΔSRCR5 and wildtype pigs. The monokine induced by gamma interferon (MIG, also known as CXCL9) was found to show consistently higher levels in wildtype pigs until day 14, when no significant difference was detected anymore. MIG is a T-cell chemoattractant to inflammation sites and involved in repair of tissue damage. In wildtype animals MIG was strongly upregulated on days 7 and 10 of the challenge (80) (
Otherwise we could see a sequence of cytokine response, with early increase of interferon α (IFNα) and interleukin-17A (IL-17A), and the interleukin 1 receptor antagonist (IL-1ra) (
Swine Health Prod 2: 10-28.
Swine Dis Comm Meet Livest Consery Inst Denver, Colo.: 173-175.
mechanisms, evolution and human disease. Nat Rev Genet 8: 762-775.
Nucleic Acid Sequences:
CD163 Guide Sequences:
Genomic Sequence of the CD163 Gene Locus in Large White Pigs (SEQ ID NO 1)
Bold=exons
Single underlined and dashed underline=splice acceptor site predictions
Double underlined=splice donor site predictions
sgRNA binding locations and cutting sites are indicated in lowercase italics, and the particular sgRNA binding to the sites is also indicated.
TTCTCAGTGC CTGCTTGGTC ACTAGTTCTC TTGGTGAGTA CTTTGACAAA TTTACTTGTA
GTGGAGGTGA AAGTGCAGGA GGAGTGGGGA ACTGTGTGTA ATAATGGCTG GGACATGGAT
GTGGTCTCTG TTGTTTGTAG GCAGCTGGGA TGTCCAACTG CTATCAAAGC CACTGGATGG
GCTAATTTTA GTGCAGGTTC TGGACGCATT TGGATGGATC ATGTTTCTTG TCGAGGGAAT
GAGTCAGCTC TCTGGGACTG CAAACATGAT GGATGGGGAA AGCATAACTG TACTCACCAA
CAGGATGCTG GAGTAACCTG CTCAGGTAAG ACATACACAA ATAAGTCAAG CCTATACATG
AACCGGTGCT TAGGAAGAAT AGAAGTCAAA TTTCAAGGAC GGTGGGGAAC AGTGTGTGAT
GATAACTTCA ACATAAATCA TGCTTCTGTG GTTTGTAAAC AACTTGAATG TGGAAGTGCT
GTCAGTTTCT CTGGTTCAGC TAATTTTGGA GAAGGTTCTG GACCAATCTG GTTTGATGAT
CTTGTATGCA ATGGAAATGA GTCAGCTCTC TGGAACTGCA AACATGAAGG ATGGGGAAAG
CACAATTGCG ATCATGCTGA GGATGCTGGA GTGATTTGCT TAAGTAAGGA CTGACCTGGG
ATGTTCAGGA AGATTGGAAG TGAAATTCCA AGGAGAATGG GGAACAATCT GTGATGATGG
CTGGGATAGT GATGATGCCG CTGTGGCATG TAAGCAACTG GGATGTCCAA CTGCTGTCAC
TGCCATTGGT CGAGTTAACG CCAGTGAGGG AACTGGACAC ATTTGGCTTG ACAGTGTTTC
TTGCCATGGA CACGAGTCTG CTCTCTGGCA GTGTAGACAC CATGAATGGG GAAAGCATTA
TTGCAATCAT AATGAAGATG CTGGTGTGAC ATGTTCTGGT AAGTGAAAAC AAAACACCGG
GGAAATTCAG AAACTGGTAG GAAAAGTGTG TGATAGAAGC TGGGGACTGA AAGAAGCTGA
TGTGGTTTGC AGGCAGCTGG GATGTGGATC TGCACTCAAA ACATCATATC AAGTTTATTC
CAAAACCAAG GCAACAAACA CATGGCTGTT TGTAAGCAGC TGTAATGGAA ATGAAACTTC
TCTTTGGGAC TGCAAGAATT GGCAGTGGGG TGGACTTAGT TGTGATCACT ATGACGAAGC
CAAAATTACC TGCTCAGGTA AGAATTTCAA TCAATGTGTT AGGAAATTGC ATTCTACTTT
ACACGTGGGG CACCGTCTGT GATTCTGACT TCTCTCTGGA GGCGGCCAGC GTGCTGTGCA
GGGAACTACA GTGCGGCACT GTGGTTTCCC TCCTGGGGGG AGCTCACTTT GGAGAAGGAA
GTGGACAGAT CTGGGCTGAA GAATTCCAGT GTGAGGGGCA CGAGTCCCAC CTTTCACTCT
GCCCAGTAGC ACCCCGCCCT GACGGGACAT GTAGCCACAG CAGGGACGTC GGCGTAGTC
T
GCTCAAGTGA GACCCAGGGA ATGTGTTCAC TTTGTTccca tgccatgaag agggtaGGGT
GGGTCCCTCT GCAACTCTCA CTGGGACATG GAAGATGCCC ATGTTTTATG CCAGCAGCTT
AAATGTGGAG TTGCCCTTTC TATCCCGGGA GGAGCACCTT TTGGGAAAGG AAGTGAGCAG
GTCTGGAGGC ACATGTTTCA CTGCACTGGG ACTGAGAAGC ACATGGGAGA TTGTTCCGTC
ACTGCTCTGG GCGCATCACT CTGTTCTTCA GGGCAAGTGG CCTCTGTAAT C
TGCTCAGGT
AAGAGAATAA GGGCAGCCAG TGATGAGCCA CTCATGACGG TGCCTTAAGA GTGGGTGTAC
GTCAGACACT ATCCCCGTGC AATTCATCAT CCTCGGACCC ATCAAGCTCT ATTATTTCAG
AAGAAAATGG TGTTGCCTGC ATAGGTGAGA ATCAGTGACC AACCTATGAA AATGATCTCA
GCCTGGTCGA TGGAGGTGGT CGTTGTGCTG GGAGAGTAGA GGTCTATCAT GAGGGCTCCT
GGGGCACCAT CTGTGATGAC AGCTGGGACC TGAATGATGC CCATGTGGTG TGCAAACAGC
TGAGCTGTGG ATGGGCCATT AATGCCACTG GTTCTGCTCA TTTTGGGGAA GGAACAGGGC
CCATTTGGCT GGATGAGATA AACTGTAATG GAAAAGAATC TCATATTTGG CAATGCCACT
CACATGGTTG GGGGCGGCAC AATTGCAGGC ATAAGGAGGA TGCAGGAGTC ATCTGCTCGG
TGATCAGTGA AAACAGCAGA GAGACCTGTG CAGGGCGCCT GGAAGTTTTT TACAACGGAG
CTTGGGGCAG CGTTGGCAGG AATAGCATGT CTCCAGCCAC AGTGGGGGTG GTATGCAGGC
AGCTGGGCTG TGCAGACAGA GGGGACATCA GCCCTGCATC TTCAGACAAG ACAGTGTCCA
GGCACATGTG GGTGGACAAT GTTCAGTGTC CTAAAGGACC TGACACCCTA TGGCAGTGCC
CATCATCTCC ATGGAAGAAG AGACTGGCCA GCCCCTCAGA GGAGACATGG ATCACATGTG
CCAGTGAGTA TCCATTCTTT AGCGCCACTG TTATCTTCTG ATCTACCTAA GCAGAAGTGT
GGAGGTTCCT GGGGCACTGT GTGTGACGAC TCCTGGGACC TTGAAGATGC TCAGGTGGTG
TGCCGACAGC TGGGCTGTGG CTCAGCTTTG GAGGCAGGAA AAGAGGCCGC ATTTGGCCAG
GGGACTGGGC CCATATGGCT CAATGAAGTG AAGTGCAAGG GGAATGAAAC CTCCTTGTGG
GATTGTCCTG CCAGATCCTG GGGCCACAGT GACTGTGGAC ACAAGGAGGA TGCTGCTGTG
ACGTGCTCAG GTGAGGGCAG AGAGTCTGGA TTGAGCTTGG AAGCTCTGGC AGCAAAGAGA
TCTCATCGCA TTCCTCATTT GGACTCAGAA GCGAAGACAG AGGCAGCGGC TCTCAGGTCT
AGGAGGAGAG AATTCTGTCC ATCAAATTCA ATACCGGGAG ATGAATTCTT GCCTGAAAGC
AGATGAAACG GATATGCTAA ATCCCTCAGG TCCGTGGGTT CTTTGAGGGC CTGTAGCCCT
TGAAGTACAA TGAAAAGGAA AATGGGAATT ATAACCTGGT GAGGTGAGTA AAAAGAATTT
| Number | Date | Country | Kind |
|---|---|---|---|
| 1617559.8 | Oct 2016 | GB | national |
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/EP2017/076460 | 10/17/2017 | WO | 00 |
