Senescence-induced cellular reprogramming drives cnidarian whole-body regeneration

Cell fate stability is essential to maintaining ‘law and order’ in complex animals. However, high stability comes at the cost of reduced plasticity and, by extension, poor regenerative ability. This evolutionary trade-off has resulted in most modern animals being rather simple and regenerative or complex and non-regenerative. The mechanisms mediating cellular plasticity and allowing for regeneration remain unknown. We show that signals emitted by senescent cells can destabilize the differentiated state of neighboring somatic cells, reprogramming them into stem cells that are capable of driving whole-body regeneration in the cnidarian Hydractinia symbiolongicarpus. Pharmacological or genetic inhibition of senescence prevented reprogramming and regeneration. Conversely, induction of transient ectopic senescence in a regenerative context resulted in supernumerary stem cells and faster regeneration. We propose that senescence signaling is an ancient mechanism mediating cellular plasticity. Understanding the senescence environment that promotes cellular reprogramming could provide a new avenue to enhance regeneration.


INTRODUCTION
Regenerative abilities of animals are irregularly scattered among taxa but are largely reverse-correlated with their structural complexity. With some exceptions, simple animals such as sponges, cnidarians, and planarians can regenerate whole-bodies from tissue fragments while complex animals such as vertebrates have more limited capabilities to restore lost body parts (Bely and Nyberg, 2010). Regeneration requires a certain level of growth plasticity that can be realized by a resident pool of stem cells or by mechanisms that induce reprogramming of differentiated cells to provide proliferative progenitors (Tanaka and Reddien, 2011). However, the high plasticity that allows for whole-body regeneration may also compromise the integrity of complex structures and increase malignancy risk (Sánchez Alvarado and Yamanaka, 2014;Slack, 2017). Therefore, high plasticity and structural complexity rarely co-occur in one species.
A major question arising from this line of arguments concerns the nature of the mechanisms that mediate cellular plasticity in regenerative animals. If regeneration was a primitive trait in metazoans (Bely and Nyberg, 2010), these mechanisms have been lost or became ineffective in non-regenerative taxa. Therefore, studying animals with high regenerative abilities could provide insight into the lack of regeneration in other animals. In the long term, this information might be harnessed to induce plasticity in complex, non-regenerative animals, enhancing their regenerative abilities under controlled conditions. Studying regeneration in a cnidarian, we show that senescence signaling can drive somatic cell reprogramming in a regenerative context. Our results, combined with data from other animals, suggest that senescence is an ancient mechanism in animals, used to convey a stress signal to surrounding tissues, thereby driving a regenerative response. We propose that the ability to respond to a senescence signal is one of the factors underlying differential regenerative abilities in modern metazoans.

Stem cell-less tissues can regenerate whole animals that include stem cells
The cnidarian Hydractinia symbiolongicarpus -a relative of jellyfishes and coralsis a highly regenerative animal that is able to regrow a lost head within 3 days post amputation (dpa) (Bradshaw et al., 2015). Hydractinia head regeneration is driven by a population of adult migratory stem cells, known as i-cells (Gahan et al., 2016), that are normally restricted to the lower body column of the animal and can differentiate into both somatic cells and gametes . These i-cells, which can easily be visualized via Piwi1 expression, migrate to the injury site post-amputation to restore the head. However, the heads of uninjured animals are devoid of i-cells ( Figure 1A) (Bradshaw et al., 2015), so the head alone would not be expected to be able to regenerate a new body due to its lack of i-cells. Surprisingly, we have discovered that the amputated oral tips of heads (known as hypostomes) can indeed regenerate into a fully functional animal ( Figure 1B) that contains i-cells despite having no i-cells immediately post-amputation ( Figure 1C). We termed the i-cells that appeared de novo in regenerating hypostomes 'secondary i-cells', as opposed to primary i-cells that are generated during embryogenesis.
To visualize the process of secondary i-cell appearance in vivo, we generated a Piwi1 transgenic reporter animal that expressed a timer mCherry protein (Subach et al., 2009) and membrane GFP under the control of the Piwi1 genomic control elements (Bradshaw et al., 2015). While the immature timer protein could not be observed under our microscopes due to the lack of a commercial filter, i-cells expressing a bright green membrane GFP could be viewed in the live translucent animal, while mature red timer protein was readily visible in differentiated cells ( Figure 1D). The reporter transgene is silenced during differentiation, resulting in the early progeny of i-cells becoming red due to maturation of the timer protein and dim green due to the GFP's half-life ( Figure 1D). Hypostomes were then isolated from transgenic Piwi1 reporter animals and examined under a fluorescence microscope to exclude the presence of any bright green i-cells; these hypostomes were then incubated in seawater ( Figure 1D). Consistent with the experiments described above, GFP + i-cells reappeared, de novo, at 6 dpa in an otherwise mature red mCherry + background ( Figure 1D). Hence, we concluded that, following amputation, a yet unknown mechanism induces the reprogramming of differentiated somatic cells back to Piwi1 + i-cells that then drive whole body regeneration.
Based on EdU incorporation, we established that a wave of DNA synthesis occurs between 3-4 dpa ( Figure 2A). Secondary Piwi1 + i-cells appeared in the hypostome tissue around 6 dpa ( Figure 1). These cells had residual EdU signals, showing that they were derived from cells that were in S-phase two days earlier (Figures 2B and S1A) Treating hypostomes with hydroxyurea to arrest cells in S-phase did not completely abolish the upregulation of Piwi1 in some cells ( Figure 2C). However, the level of Piwi1 expression in these cells was low as compared to i-cells assessed using immunofluorescence. Moreover, hydroxyurea-treated hypostomes did not regenerate and subsequently died, showing that proliferation occurring before secondary i-cell appearance is essential for complete reprogramming of somatic cells to fully functional i-cells.
To verify that somatic cells undergo reprogramming to give rise to secondary i-cells, we amputated hypostomes from a b-tubulin::FastFT transgenic reporter animal.
These animals express mCherry in all differentiated cells but not in i-cells (DuBuc et al., 2020). We allowed the hypostomes to develop secondary i-cells, then fixed and stained them with antibodies raised against Piwi1 and mCherry. We found cells that were double-positive, showing that cells that had expressed mCherry in the transgenic reporter animal (i.e., differentiated cells) reprogrammed into Piwi1 + i-cells ( Figure 2D).
The Hydractinia main body axis is generated and maintained by Wnt/b-catenin signaling (Duffy et al., 2010;Plickert et al., 2006). We used a Wnt3 transgenic reporter animal that expresses GFP in the oral-most tip of the hypostome (DuBuc et al., 2020) to dynamically follow the polarity of the isolated hypostomes. We found that GFP expression in the reporter animal lost its oral focus within 2-3 dpa, spreading around the tissue (Figures 2E and S1B). We repeated the experiment with an Rfamide transgenic reporter animal that expresses GFP in a stereotypic fashion in the nervous system of the head (Gahan et al., 2016). As with the Wnt3 reporter, within 2-3 dpa, the GFP + neurons lost their typical oral orientation and became disorganized ( Figure 2F and S1B). We concluded that axis polarity is lost in isolated hypostomes within 3-4 dpa. Following secondary i-cell appearance, the hypostomes elongated and regained polarity, as visualized by Wnt3 and Rfamide expression in transgenic reporter animals (Figures 2E-F; 6 dpa), developing into intact yet small animals.

Senescence precedes regenerative reprogramming
We then aimed to identify the nature of the signal that induces the appearance of secondary i-cells in isolated hypostomes that were initially devoid of i-cells. We hypothesized that this signal is present in regenerating tissues a few days before both reprogramming and the emergence of secondary i-cells. Therefore, we extracted RNA from amputated hypostomes 0, 1, 3, and 6 dpa and sequenced their transcriptomes. Comparing the gene expression of each stage to its preceding one, we performed KEGG pathway analyses (Kanehisa et al., 2021) and identified cellular processes consistent with a senescence episode in 1 dpa hypostomes ( Figure 2G; Senescence is a form of irreversible cell cycle arrest in animals, induced by stress, damage, oncogene expression, or telomere attrition (Rhinn et al., 2019;Roy et al., 2020). Senescent cells secrete a cocktail of factors, collectively known as the senescence-associated secretory phenotype (SASP) (Krtolica et al., 2001); these factors induce inflammation and enhance senescence and malignancy in neighboring cells (Krtolica et al., 2001;Paramos-de-Carvalho et al., 2021). Longterm accumulation of senescent cells in tissues is thought to contribute to organismal aging (Baker et al., 2011). Conversely, a role for senescence (and short-term senescence in particular) in cellular plasticity has been reported in mammals and other vertebrates (Da Silva-Alvarez et al., 2020;Paramos-de-Carvalho et al., 2021;Ritschka et al., 2017;Walters and Yun, 2020). However, its natural context of action is not well-understood. We hypothesized that amputation injury induces senescence in some cells that then emit a signal (through SASP) that promotes reprogramming of neighboring somatic cells. This would, in turn, produce stem cells that could initiate a regenerative process.
Following this line of reasoning, we looked for indicators of senescence in regenerating hypostomes. Cell cycle regulators such as p21 (encoded by CDKN1A) and p16 (encoded by CDKN2A) are known senescence markers in mammals (Hernandez-Segura et al., 2018). The Hydractinia genome encodes three CDKN1Alike genes ( Figure S2) but no CDKN2A, the latter being vertebrate-specific. Our phylogeny could not resolve orthology between cnidarian and bilaterian CDKN1A proteins, suggesting that the Hydractinia genes were paralogs. Therefore, we called the Hydractinia three CDKN1A-like genes Cyclin-dependent kinase inhibitor 1 (Cdki1), Cdki2, and Cdki3, respectively. Single-molecule fluorescence mRNA in situ hybridization showed that the three genes were expressed in some cells along the body column (Figures 3A and S3A) but only Cdki1 was upregulated at the cut side of isolated hypostomes around 1 dpa ( Figure 3B), being nearly undetectable in intact hypostomes ( Figure 3A). Expression patterns of Cdki2 and Cdki3 did not visibly change following injury. An additional senescence indicator is senescence- In the closely related cnidarian Hydra, it has been shown that bisection induces apoptosis in i-cells, with these dying cells emitting a Wnt3 signal that drives proliferation and head regeneration (Chera et al., 2009). However, in Hydractinia regenerating hypostomes, no evidence for apoptosis was found using TUNEL staining ( Figures 3D, S4B, and S4C), suggesting that senescence-induced regeneration in the absence of stem cells is distinct from apoptosis-induced repair where i-cells are present.
Given that senescent cells do not spontaneously die or normally exit senescence, the disappearance of senescence markers between the second and third dpa prompted us to investigate their fates. For this, we generated a Cdki1 transgenic reporter animal that expressed membrane GFP CAAX under the Cdki1 genomic control  Figure   3C). Therefore, whole-body regeneration from isolated hypostomes is accompanied by a short senescence period around 1 dpa, loss of tissue polarity within 3 dpa, a burst of proliferation around 3-4 dpa, and secondary i-cell appearance 6 dpa ( Figure   4A). The process is completed by the re-establishment of tissue polarity, elongation, and morphogenesis.

Senescence signaling is required and sufficient to induce reprogramming
To identify a functional link between the short period of senescence and the subsequent reprogramming of somatic cells, we used navitoclax, a senolytic drug (Chang et al., 2016), to inhibit senescence and study the effect of this manipulation on somatic cell reprogramming and secondary i-cell appearance. We exposed freshly amputated hypostomes to 1 µM navitoclax in seawater and followed them over 6 dpa. We found that navitoclax at this concentration inhibited senescence marker upregulation for the duration of treatment ( Figure 4B). Furthermore, no secondary i-cells appeared in treated hypostomes at 6 dpa ( Figure 4C). We repeated the experiments with rapamycin, an mTOR inhibitor, which yielded similar results.
To further address the requirement of senescence to reprogramming and exclude an unspecific effect of senolytic drugs, we used CRISPR-Cas9 to mutate the Cdki1 gene ( Figure S5A). Three short guide RNAs (sgRNAs) targeting the nuclear localization signal (NLS) and PCNA-interacting domain of Cdki1 were designed and injected with recombinant Cas9 into fertilized eggs (Gahan et al., 2017). G0 embryos were allowed to develop into larvae, metamorphose, and grow to the young colony stage. They were then screened for mutations by genomic PCR and sequencing.
Confirmed mosaic mutants were grown to sexual maturity and crossed with wild type animals. Heterozygous G1 animals were identified by PCR and sequencing, grown to sexual maturity, and interbred to give rise to G2 homozygous knockout (KO) animals ( Figure S5A). These animals developed normally to the larval stage, metamorphosed, and grew to apparently normal colonies that were able to regenerate amputated heads similar to those seen in wild type animals ( Figure S5B). We amputated hypostomes from Cdki1 KO animals and analyzed their behavior. We found that the absence of functional Cdki1 resulted in the loss of the senescence response at 1 dpa ( Figure 4B) and absence of secondary i-cells at 6 dpa ( Figure   4C). With the lack of i-cells, the KO hypostomes had not regenerated within 20 dpa ( Figure 4D), remaining as amorphous tissue lumps that eventually died of starvation. Therefore, a short senescence signaling is essential for reprogramming.

However, EdU analysis revealed that
Finally, we tested the ability of ectopic senescence to induce reprogramming in this specific context. To induce senescence, we employed an optogenetic approach using the Opto-SOS genetic cassette (Johnson et al., 2017). Cells carrying this construct respond to blue light by overactivation of the Ras pathway ( Figure 5A), and overactivation of Ras is known to induce senescence (Serrano et al., 1997). We generated transgenic mosaic animals that carried the Opto-SOS construct, fused to mScarlet. Hypostomes were amputated and exposed to blue light for 12 hpa ( Figure   5B) and we then observed the events accompanying their regeneration. As expected, we found that exposure to blue light induced enhanced SAb-Gal activity ( Figures 5C-D), and secondary i-cells appeared 5 dpa as opposed to 6 dpa in animals kept in the dark. At 6 dpa, the number of secondary i-cells was significantly enhanced in animals exposed to blue light comparing to those kept in the dark presented here are consistent with the notion that senescence signaling is one of the factors that mediate cellular plasticity (Rhinn et al., 2019). However, the ability to respond to a senescence signal has not been highly conserved across lineages during metazoan evolution.
A rudimentary response to senescence signals by increased plasticity is still present in modern mammals. This has been shown in mouse liver cells (Ritschka et al., 2017), skeletal muscles (Chiche et al., 2017), and by the discovery that a senescent environment facilitates reprogramming by OSKM factors (Mosteiro et al., 2016).
Finally, embryonic and induced pluripotent stem cells maintain pluripotency when grown on a feeder layer consisting of senescent fibroblasts (Takahashi and Yamanaka, 2006). However, except for urodele amphibians (Walters and Yun, 2020), tetrapod vertebrates have poor regenerative ability and do not respond to a senescence signal as effectively as Hydractinia.
We suggest that senescence is an ancient mechanism, instructing cells adjacent to an injury site to prepare for a regenerative event. We also speculate that other consequences of senescence that have been observed in mammals, such as longterm retention and accumulation of senescent cells, aging, chronic inflammation, and cancer, are side effects that evolved later in the evolution of these lineages, perhaps as consequence of the increase in cell fate stability and morphological complexity.
Understanding the senescent environment and its role in cellular plasticity could pave the way for new treatments to enhance regeneration in poorly regenerating mammals.

Limitations of the study
Our study provides strong evidence for a role for senescence signaling in cellular reprograming in cnidarians. While evidence from the literature provides indications to the existence of similar phenomena in other animals, the degree to which components of the senescence signaling pathway are evolutionarily conserved across phyla is at present unclear. Furthermore, markers for cellular senescence are not universal and no definitive senescence marker has been identified. Finally, the existence of different senescence 'types' has been proposed (Varela-Eirin and Demaria, 2022). Future work on other metazoans will be required to establish the pan-animal role of senescence signaling.

Author contribution
MSS and UF conceptualized the study. MSS performed experiments. GK performed phylogeny and Tunel assays. HRH and F analyzed data. ADB supervised the generation of RNA-seq data and making these data publicly available as described below. MSS and UF designed the experiments, analyzed data, and wrote the paper.
All authors commented on and approved the manuscript.

Data availability
The Hydractinia symbiolongicarpus genome is available in the Hydractinia Genome Project Portal through the NIH National Human Genome Research Institute       CDKN-like, mTor, and p53-like genes. (A-B), Topology of metazoan CDKN phylogeny obtained by Bayesian inference (A) and Maximum likelihood (B). Topologies show high divergence between CDKN sequences, numerous unresolved nodes, and inconsistency with species phylogeny. The three Hydractinia CDKI1, 2, and 3 named to indicate lack of clear orthologous relationships with vertebrate homologues. (C-D), Topology of metazoan mTOR phylogeny obtained by Bayesian inference (C) and Maximum likelihood (D). The topologies are consistent with metazoans phylogeny: cnidarian sequences are monophyletics and the clade is sister-group to bilaterians. Inside bilaterians, two strongly supported monophyletic groups are identified, the deuterostomian and protostome genes (ecdysozoans+molluscs). Similarity between species phylogeny and mTOR phylogeny reflect the conservation of this gene during evolution. (E-F), Topology of metazoan P53-like family phylogeny obtained by Bayesian inference (E) and Maximum likelihood (F). Bilaterian sequences are not monophyletic in Bayesian analysis, probably due to Long Branch attraction of ecdysozoan genes grouped with few cnidarian sequences. Most cnidarian sequences are distributed close to the root. In Maximum likelihood analysis, bilaterian genes are monophyletic and present coherent topology according to species phylogeny: deuterostomes are sister-group to protostomes (ecdysozoans+molluscs). P53, P63, and P73 are vertebrate specific paralogs. Consequently, the two Hydractinia P53 are named P53 like 1 and 2.

Hydractinia husbandry
Adult Hydractinia symbiolongicarpus colonies were maintained as described in ref (Frank et al., 2020). Female and male colonies were grown on glass slides kept in artificial seawater (ASW) at 19-22ºC. Animals were fed four times per week with Artemia nauplii, and once a week with pureed oysters. To induce scheduled spawning, we kept the animals in a constant 14:10 light:dark cycle, where females and males spawn 1.5 hours after exposure to light.

Hypostome isolation
One day starved adult H. symbiolongicarpus colonies were anaesthetized in 4% MgCl2 (in 50% distilled water / 50% filtered seawater). Polyps were dissected from the colony and decapitated. Hypostomes were isolated by removing the tentacles. Hypostomes were transferred to a glass Petri dish with freshly 0.2 µm filtered seawater and incubated at 22.4-23˚C with constant reciprocal shaking in a temperature-controlled incubator. 0.2 µm filtered sea water was changed every two days.

RNA Isolation from hypostome tissue
Total RNA was extracted from isolated hypostomes collected from wild type H. symbiolongicarpus strain 291-10. Samples consisted of three replicates of 220 hypostomes each at 0-, 1-, 3-, and 6-days post amputation (dpa). RNA was isolated using Direct-zol TM RNA MiniPrep kit (Zymo Research; R2050) per the manufacturer's instructions. The eluted RNA was quantified using NanoDrop and the quality was checked using agarose-formaldehyde gel electrophoresis. After RNA isolation, samples were shipped to the NIH Intramural sequencing (NISC) for further processing and sequencing. RNA was amplified with the Ovation RNA-Seq System V2 kit, and sequencing libraries were made with Illumina TruSeq Stranded mRNA Library Prep Kit. Libraries were sequenced on one lane of Illumina NovaSeq 6000 (2x151 bp), generating between 87 and 137 million reads per sample (average 112 million reads). Only two replicates for 6 dpa were sequenced.

Differential expression and gene set enrichment analysis
Raw reads were trimmed and aligned to the reference genome using Trim Galore https://www.bioinformatics.babraham.ac.uk/projects/trim_galore/ and HISAT2 (Kim et al., 2015), respectively. A count matrix of mapped reads per genomic feature was generated using Subread featureCounts (Liao et al., 2014) and converted into a DESeqDataSet. This dataset was used as an input to DESeq2 (Love et al., 2014) in order to generate a list of differentially expressed genes for each time point compared to the subsequent time point. Lists of differentially expressed genes were then separated based on up vs down regulated genes; sorted from the highest log2fold changes to the lowest. These lists then used as an input for gene set enrichment analysis. To assign KEGG_Pathway terms to each gene ID, we fed Hydractinia symbiolongicarpus protein sequences, as coded in the recent genome (https://research.nhgri.nih.gov/hydractinia/download/protein_models/symbio/Hsym_p rimary_v1.0.aa.gz), into EggNOG-Mapper v5.0 (Huerta-Cepas et al., 2019), which then used as reference in ClusterProfiler. Gene set enrichment analysis were performed by using lists of differentially expressed genes as input into ClusterProfiler::enricher (Wu et al., 2021). We then plotted the identified enriched pathways (adj.p-value < 0.05) in Microsoft Excel.

Transgenic animals
The generation of stable transgenic reporter animals was carried out as previously described . Wnt3::GFP (DuBuc et al., 2020) and RFamide::GFP  reporters were previously generated in our lab. We used the GENEius online tool from Eurofins Genomics to codon optimize all the synthetic gene sequences for Hydractinia symbiolongicarpus. All synthetic genes were synthetized using IDT gBlocks gene fragments. Vector maps can be found below.
Fast Fluorescent timer protein reporter (FastFT): The codon optimized FastFT coding sequence (Subach et al., 2009) was designed in frame with P2A peptide and membrane GFP sequences (FastFT-P2A-GFP CAAX ). The synthetic sequence was amplified by PCR and inserted into the Piwi1 reporter plasmid (Bradshaw et al., 2015) using NotI and SacI restriction enzymes. G0 colonies were bred to G2 (nonmosaic transgenic offspring). The timer property of the FastFT fluorescent protein (earlier blue and later red fluorescent forms according to maturation state) was aimed to differentiate i-cells (recently translated protein) from their progeny. The mature red form was detected using an mCherry filter set. However, our microscopes lacked the proper filter setup to detect the FastFT blue form. Thus, to identify i-cells, we calibrated the laser power to higher GFP intensities (DuBuc et al., 2020) that were devoid of or low in mCherry fluorescence (FastFT mature red form). Cdki1 reporter: 5' upstream and 3' downstream regulatory sequences of the Cdki1 gene (HyS0010.253) were cloned from extracted genomic DNA by PCR and inserted into an open cloning vector. Membrane GFP sequence (GFP CAAX ) was placed in frame with the 5' upstream regulatory sequence. G0 colonies were bred to G2 (nonmosaic transgenic offspring). OptoSOS: The OptoSos coding sequence (Johnson et al., 2017) was codon optimized and designed replacing the RFP sequence by our codon optimized mScarlet fluorescent protein . The synthetic sequence was amplified by PCR and inserted into the ß-tubulin reporter construct  using NotI and SacI restriction enzymes. We used BLAST to identify the endogenous H. symbiolongicarpus SOS catalytic domain that was amplified by PCR and inserted in frame with the SSPB sequence. Single injected embryos were grown to adult mosaic colonies, and we selected the animals who expressed the transgene in the hypostome.

Single polyp genomic DNA extraction
Protocol was adapted from ref . A single polyp was isolated from a colony and transferred to 1.5 ml Eppendorf tube. The polyp was suspended in 50 µL Lysis buffer (10 mM Tris pH 8.0, 10 mM EDTA, 2% SDS), flicked few times, supplemented with 50 µL of digestion buffer (10% SDS, 0.4 mg/mL Proteinase K), and incubated at 56˚C for 2-3 hours with occasional flicking. Then, 100 µL of cold phenol-chloroform (pH 8.0) was added. Samples were centrifuged at maximum speed for 15 minutes at 4˚C, and supernatant was transferred to a new tube. Genomic DNA precipitation was carried out overnight at -20˚C by adding two volumes of isopropanol and 1:10 total volume of Sodium Acetate Solution (3 M). Samples were cleaned up at 4˚C by centrifugation and a 70% Ethanol wash. Extracted genomic DNA was dried out, resuspended in 10 µL nuclease-free water, and stored for further genotyping PCR.

Genotyping CRISPR/Cas9 knock-outs
One hundred CRISPR-Cas9 larvae that had been injected as zygotes were metamorphosed and grown into small colonies. Of these, forty animals were analyzed for Cdki1 mutations. Primers spanning the entire coding region of the gene were used in a PCR reaction to identify large deletions. We kept and grew eight G0 mosaic colonies to sexual maturity and crossed one male with a wild-type female. One hundred G1 animals were metamorphosed, grown to small colonies, and genotyped. Two sets of primers were used to identify heterozygous colonies carrying the genomic deletion: one spanning the entire coding region and another internal primer targeting the deleted intron. PCR products of heterozygous colonies were cloned into PGEMT-easy vector and sequenced using the T7 and SP6 primers. Each of these animals was a heterozygote carrying a 3' deletion mutation in one Cdki1 allele at the predicted cut sites and one wild type allele. One male and one female were crossed and G2 offspring were genotyped identifying five mutant homozygous animals. We performed genotyping on these animals from two independent DNA extractions and confirmed the mutation by sequencing.