Loss of the homologous recombination gene rad51leads to Fanconi anemia-like symptoms in zebrafishJan Gregor Botthofa,b,c, Ewa Bielczyk-Maczy�nskaa,b,d,1, Lauren Ferreiraa,b,c, and Ana Cvejica,b,c,2

aDepartment of Haematology, University of Cambridge, Addenbrookes Hospital, Cambridge CB2 0XY, United Kingdom; bWellcome Trust Sanger Institute,Cambridge CB10 1SA, United Kingdom; cWellcome Trust–Medical Research Council Cambridge Stem Cell Institute, Cambridge CB2 1QR, United Kingdom;and dNational Health Service Blood and Transplant, Cambridge CB2 0PT, United Kingdom

Edited by Stephen C. Kowalczykowski, University of California, Davis, CA, and approved April 25, 2017 (received for review December 19, 2016)

RAD51 is an indispensable homologous recombination protein, nec-essary for strand invasion and crossing over. It has recently beendesignated as a Fanconi anemia (FA) gene, following the discoveryof two patients carrying dominant-negative mutations. FA is a hered-itary DNA-repair disorder characterized by various congenital abnor-malities, progressive bone marrow failure, and cancer predisposition.In this report, we describe a viable vertebrate model of RAD51 loss.Zebrafish rad51 loss-of-function mutants developed key features ofFA, including hypocellular kidney marrow, sensitivity to cross-linkingagents, and decreased size. We show that some of these symptomsstem from both decreased proliferation and increased apoptosis ofembryonic hematopoietic stem and progenitor cells. Comutation ofp53 was able to rescue the hematopoietic defects seen in the singlemutants, but led to tumor development. We further demonstratethat prolonged inflammatory stress can exacerbate the hematologi-cal impairment, leading to an additional decrease in kidney marrowcell numbers. These findings strengthen the assignment of RAD51 asa Fanconi gene and provide more evidence for the notion that aber-rant p53 signaling during embryogenesis leads to the hematologicaldefects seen later in life in FA. Further research on this zebrafish FAmodel will lead to a deeper understanding of the molecular basis ofbone marrow failure in FA and the cellular role of RAD51.

Fanconi anemia | stem cells | hematopoiesis | cytokine effects |inflammation

Fanconi anemia (FA) is a hereditary DNA-repair disorder char-acterized by various congenital abnormalities, progressive

bone marrow failure (BMF), and cancer predisposition (1). It iscaused by mutations in one of 21 genes in the FA pathway (2, 3)(www2.rockefeller.edu/fanconi/). The FA pathway has beenshown to be the major route for the removal of interstrand cross-links (ICL): DNA lesions that prevent replication and transcriptionby inhibiting DNA strand separation (4, 5). When the pathway isdefective, these structures cannot be removed, potentially leadingto cell death (6). Indeed, sensitivity to cross-linking agents, such asmitomycin C (MMC), is an absolute diagnostic criterion of FA (7).Although FA is characterized by remarkable phenotypic hetero-

geneity, FA patients usually succumb to the depletion of hemato-poietic stem and progenitor cells (HSPCs) in their BM, leading topancytopenia and complete BMF. Therefore, BM transplantation isthe only modality that offers a potential cure of hematopoietic de-fects but is itself associated with considerable morbidity (8, 9). In-terestingly, a decrease in HSPCs (CD34+ cells) is already apparentin FA infants even before the first hematological symptoms appear(10). This finding led to the hypothesis that FA originates fromdefects during the formation of the initial HSPC pool, presumablybecause of an overactive p53/p21 response and cell cycle arrest (10).In agreement with this, FA mice have considerably smaller fetallivers than their healthy siblings (11). It remains unclear, however, atwhich stage during embryonic development these defects appearand how perturbation in the production of embryonic HSPCs relatesto the phenotype seen in adulthood.Because of the role FA genes play in the repair of ICLs, DNA

damaging agents causing ICLs have been proposed as a major

cause of BMF, with small aldehydes being the most likely candi-dates. Comutation of genes in the FA pathway and aldehydemetabolizing genes (Aldh2 and Adh5) resulted in significant re-duction of HSPCs and BMF in double-mutant mice (12–16). Inaddition, FA patients lacking ALDH2 show a more severephenotype (17, 18). Apart from their hypersensitivity to cross-linking agents, FA cells also react excessively to proapoptoticcytokines, such as IFN-γ and TNF-α (19–24). However, therole of cytokines in the etiology of BMF remains controversial(25–28).In the last 2 y, a novel FA subtype associated with dominant-

negative mutations in RAD51 has been reported, leading to thedesignation of RAD51 as FANCR (29–31). It has been shown to beinvolved in protecting broken down replication forks from excessprocessing by nucleases, linking the FA pathway with RAD51/BRCA2 (29, 32). In vivo studies of Rad51 have previously beenvery difficult, as mice lacking the protein invariably die during earlyembryogenesis (33, 34).In this study, we characterized a viable vertebrate model of

Rad51 loss. Indeed, our zebrafish rad51 loss-of-function mutantrecapitulates many congenital and hematological features ofFA. We provide in vivo evidence that decreased HSPC num-bers during embryonic development directly lead to the laterBM defects in FA. Finally, we show that rad51 mutants do notoverproduce inflammatory cytokines, but are more sensitiveto them and that prolonged inflammatory stress can furtherreduce marrow cellularity.

Significance

The homologous recombination protein RAD51 has been exten-sively studied in prokaryotes and lower eukaryotes. However,there is a significant lack of knowledge of the role of this proteinand its regulation in an in vivo context in vertebrates. Here wereport the first viable vertebrate mutant model of rad51 inzebrafish. These mutant fish enabled us to confirm the recentlydiscovered role of RAD51 in Fanconi anemia pathogenesis. Wereport that p53-linked embryonic stem cell defects directly lead tohematological impairments later in life. Comutation of rad51with p53 rescues the observed hematological defects, but pre-disposes the fish to early tumor development. The application ofthis model opens new possibilities to advance Fanconi anemiadrug discovery.

Author contributions: J.G.B., E.B.-M., and A.C. designed research; J.G.B., E.B.-M., and L.F.performed research; J.G.B., E.B.-M., and A.C. analyzed data; and J.G.B. and A.C. wrotethe paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.1Present address: Department of Chemical and Systems Biology, Stanford University,Stanford, CA 94305.

2To whom correspondence should be addressed. Email: as889@cam.ac.uk.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1620631114/-/DCSupplemental.

E4452–E4461 | PNAS | Published online May 16, 2017 www.pnas.org/cgi/doi/10.1073/pnas.1620631114

ResultsThe rad51sa23805 Allele Leads to Complete Loss of Functional Rad51.To study the function of Rad51 in hematopoiesis, we obtainedfish carrying the rad51sa23805 allele from the Sanger InstituteZebrafish Mutation Project (35). The rad51sa23805 allele has aC > T mutation at codon 203 in exon 7, which leads to a pre-mature stop codon in the region of the RecA domain. In con-trast to mice lacking Rad51, which invariably die during earlydevelopment (33, 34), fish carrying homozygous copies of therad51sa23805 (referred to as rad51−/− for brevity in the text)survive to adulthood. However, all surviving adults undergo sex

reversal and are infertile males, lacking mature spermatozoa inthe testes (SI Appendix, Fig. S1).To ensure complete loss-of-function of the rad51sa23805 allele, we

carried out a Western blot on testicular tissue, because Rad51 ishighly expressed in germ cells in zebrafish and other species (36,37). This approach confirmed that the full-length protein is lost inrad51−/− fish (Fig. 1A). However, immunostaining for Rad51 at theone-cell stage embryos (i.e., before maternal to zygotic transition)(38) showed diffuse staining in the cytoplasm of all embryosresulting from a rad51+/− in-cross (Fig. 1B). This finding suggestedthat rad51−/− embryos have maternally derived Rad51.

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Fig. 1. The rad51sa23805 allele leads to loss of Rad51 protein and causes DNA damage sensitivity. (A) Western blot showing the expression of Rad51 in testesextracts of WT and mutant zebrafish. β-Actin was used as a loading control. (B) Representative Rad51 immunostained embryos derived from a rad51+/− in-cross. Sec-ondary only (i), Abcam primary (ii), AnaSpec primary (iii). (Magnification, 65×.) (C) Chromosome spreads of 24-hpfWT andmutant embryos treated with 1 μg/mL DEB for20 h taken using a 100× oil-immersion objective. White arrows indicate characteristic damage (chromosome breaks and radial structures) in response to cross-linkingagents (i). Quantification of the damage (ii). Mann–Whitney test, P = 0.0004, n+/+ = 25, n−/− = 26. (D) Comparison of the response of 48-hpf WT and mutant embryos toirradiation (i). (Magnification, 15×.) The black arrow indicates the small head and eye phenotype, which is quantified in ii. Two-tailed Fisher’s exact test pooling WT andheterozygotes as control group, P < 0.001, n = 67. (E) Immunostaining for pH2AX in WT and mutant embryos with pictures of representative embryos taken with a 63×water-immersion objective (i) and quantification of foci (ii). White arrows indicate example foci. Two-tailed Student’s t test, P < 0.0001, n+/+ = 8, n−/− = 11.

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Because of the early embryonic lethality of murine Rad51 mu-tants (33, 34), we decided to look at the functional redundancybetween rad51 and its paralogues. To do this, we knocked outrad51l1, the fish ortholog of the human RAD51B gene usingCRISPR. Initially, we raised 204 embryos resulting from a rad51+/−,rad51l1+/− in-cross to adulthood. However, of these none weredouble mutants, even though 12.75 would be expected fromMendelian inheritance. We repeated the same cross and collectedembryos at 4 d postfertilization (dpf). Again, 0 of 44 embryos weredouble mutants, even though 2.75 would have been expected. Fi-nally, we repeated the cross and genotyped at 6 h postfertilization(hpf; the earliest point the genotyping works reliably). Of 47 em-bryos, none were double mutants. This early embryonic syntheticlethality of comutation indicates that rad51 and rad51l1 have atleast some redundant functionality.

Lack of Rad51 in Embryos Leads to Increased DNA Damage Sensitivity.Rad51 is essential for the repair of ICLs and double-strandedbreaks via homologous recombination (HR). In ICL repair, it playsa role in both HR-linked processes, as well as HR-independentsteps (29–31). To investigate the role of Rad51 in the repair ofICLs, we treated embryos resulting from an in-cross of rad51+/−

parents with the cross-linking agents diepoxybutane (DEB) andMMC, as well as the topoisomerase I inhibitor camptothecin(CPT) and the poly(ADP-ribose) polymerase (PARP)-1 inhibitor1,5-isoquinolinediol (DiQ). After treatment, the tails of the em-bryos were used for chromosome spreads, whereas the heads werekept for genotyping. The spreads revealed that DEB induced de-fects characteristic for FA, including breaks and radial structures(Fig. 1C). MMC induced many of the same features, but also led toconsiderably more premature chromatid separation events (SIAppendix, Fig. S2A). Similarly, CPT induced high levels of chro-mosome breaks, but few radial structures (SI Appendix, Fig. S2B).DiQ, on the other hand, was embryonic-lethal at higher concen-tration (100 μM), but at lower levels (10 μM) induced just a fewbreaks in both WT and mutant embryos (SI Appendix, Fig. S2C).We also examined the role of Rad51 in the repair of other forms

of double-stranded breaks not involving cross-linking agents. Forthis, we irradiated 24-h-old embryos from an in-cross of rad51+/−

parents with γ-radiation and examined them at 48 hpf. The irra-diated embryos were then blindly imaged, scored, and genotyped.The rad51−/− embryos developed small eyes and heads in responseto radiation (Fig. 1D, i). This phenotype was limited to the rad51−/−

embryos (Fig. 1 D, ii), showing that they are more sensitive to ir-radiation than WT siblings. In addition, pH2AX (a marker ofdouble-stranded breaks) immunostaining of nonirradiated em-bryos revealed extensive DNA damage in embryos lackingRad51 in comparison with their WT siblings (Fig. 1E). Takentogether, these data suggest that rad51−/− embryos are hypersen-sitive to DNA damage.Finally, we considered the role of nonhomologous end-joining

(NHEJ) in the survival of our mutant fish. To do that, we treatedan in-cross of rad51+/− parents with a range of different concen-trations of SCR-7, a DNA ligase IV inhibitor, and scored theembryos at 24 hpf (SI Appendix, Fig. S3). At the lowest concen-tration, embryos were completely unaffected, whereas at the higherconcentration all embryos died. At no concentration were themutant embryos more sensitive than their WT siblings (SI Appen-dix, Fig. S3), indicating that NHEJ is dispensable for the survival ofrad51−/− embryos.

Rad51 Mutants Recapitulate Many Congenital and HematologicalFeatures of FA. RAD51 mutations were recently linked to FA inhumans in two case reports (29, 30). Therefore, we examined therad51 mutant fish for congenital and hematological features asso-ciated with FA. A common congenital symptom of FA is decreasedheight (1). On account of that, we measured the body length of fishwith and without rad51 mutation throughout development. Al-

though embryonic development was not affected by the mutation,the size of rad51−/− fish was decreased compared with their WTsiblings, starting from around 23 dpf (SI Appendix, Fig. S4A). Thesize reduction during the larval period was maintained to adulthoodwith rad51 mutant fish being on average 10% shorter than theirWT siblings (SI Appendix, Fig. S4 B and C). In addition, rad51−/−

embryos and larvae developed microphthalmia, another distinctfeature of FA patients (SI Appendix, Fig. S5).To assess whether loss of rad51 affects adult hematopoiesis, we

inspected the kidney marrow (WKM) of zebrafish, which is anal-ogous to BM in mammals. Unlike in mammals, there is only onekidney in zebrafish, which is the only site of adult hematopoiesis.Importantly, there is no space restriction of the marrow by the rigidbone, as kidney tissue is very spongy and flexible. We first obtainedseveral H&E-stained histological sections each of the kidney offour WT and mutant fish, ranging between 4 and 8 mo of age.Representative sections showed no noticeable qualitative differ-ences between rad51+/+ and rad51−/− fish (Fig. 2A). However, eventhough the morphology appeared normal, kidney size and cellnumber were considerably (∼50%) decreased in rad51−/− fish (Fig.2B). The kidney cellularity of rad51−/− fish gradually decreasedwith aging, but at the same rate as in WT fish (Fig. 2 B, ii), sug-gesting that the mutants established steady-state hematopoiesisdespite displaying kidney hypocellularity. Although rad51 mutantsdid not develop cytopenia in the peripheral blood (PB) (Fig. 2C),they did accumulate macrocytic erythrocytes in the PB (Fig. 2D),suggesting a gradual worsening of the phenotype over time.

Rad51−/− Fish Show a Hyperproliferative Phenotype in the WKM. Thenumber of cells in the WKM is determined by the balance betweentheir proliferation and apoptosis rate, as well as their migration tothe PB. To test whether the decrease in kidney cell numbers inrad51 mutants was a result of apoptosis, we carried out an AnnexinV-propodium iodide (AV-PI) staining assay on kidney tissue (Fig.3A and SI Appendix, Fig. S6 A and B). We observed no statisticallysignificant difference between rad51+/+ and rad51−/− fish, excludingapoptosis as an initial cause for the decrease in cell numbers in thekidney. Therefore, we next focused on the proliferation of thekidney marrow cells.The standard assay for assessing cell proliferation rates is the

incorporation of BrdU into the DNA. To follow the kinetics ofBrdU incorporation and dilution via division or migration tothe circulation, we measured BrdU labeling at several timepoints postinjection (Fig. 3B). Because erythrocytes are nu-cleated in zebrafish, our analysis was not limited only to leu-kocytes and allowed us to robustly assess changes in the WKMas well as in the PB. The initial number of BrdU+ cells in theWKM was twofold higher in rad51−/− fish compared with thecontrol (Fig. 3 C, i and SI Appendix, Fig. S6F), suggesting anincreased proliferation rate in the mutant. This was followedby a fast dilution of the BrdU label over 2 wk, because of theincreased cell division of mutant HSPCs (Fig. 3 C, i). In linewith this finding, the initial number of BrdU+ cells in the PBwas higher in rad51−/− fish compared with WT siblings (Fig. 3C, ii), but then plateaued faster because of the dilution of thelabel in the kidney (Fig. 3 C, ii).To show that blood, rather than other cells in the WKM, are

proliferating, we generated mutants in various transgenic back-grounds. The rad51−/− Tg(gata1a:EGFP) line (39) was used to as-sess the erythrocytic lineage, including erythrocytic progenitors.The rad51−/− Tg(cd41/itga2b:EGFP) (40) line was used to detectthrombocytic progenitors, which are labeled in the cd41:EGFPdim

subpopulation (41). Consistent with the BrdU incorporation ex-periments, we saw an increase in newly made gata1:EGFP+

erythrocytes in the kidney (Fig. 3D and SI Appendix, Fig. S6C), aswell as an increase in cd41:EGFPdim thrombocytic progenitors (Fig.3E and SI Appendix, Fig. S6D).

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Together, this evidence shows that loss of rad51 leads to ahyperproliferation of HSPCs, possibly as a compensatory mecha-nism to prevent cytopenia in the PB. However, the lowered cel-lularity of the kidney in adult rad51−/− fish cannot be explained bythis finding, suggesting that the kidney cytopenia stems from anearly, possibly embryonic HSPC defect.

Lack of Rad51 Causes an HSPC Defect During Early Development. Thedefinitive wave of hematopoiesis starts at around 30 hpf, whenlong-term hematopoietic stem cells (HSCs) are formed fromendothelial cells of the ventral wall of the dorsal aorta. Thesenewly made HSCs move to the caudal hematopoietic tissue(CHT), which serves as an intermediate place of hematopoiesis

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Fig. 2. Adult rad51 mutant fish display kidney marrow cytopenia. (A) H&E-stained histological sections of 4-mpf WT and mutant kidneys using a 20× objective.Muscle (red arrow), ducts/tubules (gray arrow), and hematopoietic kidney marrow (black arrow) can be seen. (Scale bar, 100 μm.) (B) Fixed 8-mpf WT and mutantkidneys (i) (magnification, 10×); quantification of the number of total cells per freshly isolated kidney at different ages using a hemocytometer (ii). Two-wayANOVA was used and type III model fit [Armitage et al. (73)]. The test shows a significant influence of age [F(1, 50) = 18.23, P < 0.0001] and mutation status[F(1, 50) = 10.87, P = 0.0018] on phenotype. Fourmonths postfertilization n+/+ = 6, n−/−= 6; 8 mpf n+/+ = 16, n−/−= 16; 13 mpf n+/+ = 6, n−/−= 4. (C) Quantification ofPB cells in WT and mutant fish at 4 mpf. Two-sided t test, n+/+ = 6, n−/−= 6. n.s., not significant. (D) In i, blood smears of 13-mpfWT (Upper) and mutant fish (Lower)are compared. (Scale bar, 10 μm.) In ii, the change is quantified using two-way ANOVA and a type III model fit [Armitage et al. (73)]. There was a statisticallysignificant interaction between age and mutation status [F(1, 28) = 12.89, P = 0.0012], no significant influence of age [F(1, 28) = 180.76, P = 0.392] and nosignificant influence of mutation status [F(1, 28) = 2.88, P = 0.1006]. P value shown on the graph stems from a post hoc Tukey multiple-comparison test. Fourmonths postfertilization: n+/+ = 6, n−/− = 6; 8 mpf: n+/+ = 5, n−/− = 5; 13 mpf: n+/+ = 6, n−/− = 4. Bars represent mean ± SEM.

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in which the HSCs expand greatly, akin to the mammalian fetalliver (42).To assess the underlying cause of the decreased number of cells

in the adult kidney of FA fish, we focused on embryonic hema-topoiesis. To this end, we used whole-mount in situ hybridization(ISH) using a cmyb-specific probe, which labels HSPCs. Indeed, at

2 dpf, rad51−/− embryos had a decreased number of HSPCs com-pared with the WT embryos from the same clutch (Fig. 4A). At4 dpf, the difference in the number of HSPCs in the CHT ofrad51−/− and WT embryos was further exacerbated (Fig. 4B).Following up on that finding, we carried out a BrdU in-

corporation assay on the tail tissue of 2-dpf embryos, which showedthat the proliferation rate in the tail of rad51−/− embryos was abouthalf that of rad51+/+ embryos (Fig. 4C). Although not statisticallysignificant, this trend was still apparent in the CHT at 4 dpf(Fig. 4D).In addition to proliferation, we also investigated apoptosis at

2 dpf by carrying out a TUNEL assay on several crosses of rad51heterozygotes. This process revealed a twofold increase in apo-ptosis in the CHT in mutants compared with WT embryos (Fig. 4 Eand F). Taken together, our data imply that the cytopenia in theadult kidney is caused by an increase apoptosis of HSPCs, as wellas reduced proliferation during early embryogenesis, mainlybefore 4 dpf.

The HSPC Defects in rad51−/− Fish Are Mediated via p53. The defectsseen in FA have recently been linked to an aggravated p53 response(10). To focus on the role of p53 in the HSPC defect, we generateda zebrafish line carrying mutations in both rad51 and p53. Thisdouble mutation was able to rescue the number of HSPCs in theCHT of 4-dpf embryos (Fig. 5A and SI Appendix, Table S3). Im-portantly, the number of cells in the adult WKM of p53−/− rad51−/−

fish reached WT levels by 4 m postfertilization (mpf) (Fig. 5B).Along with the rescued marrow cellularity, there was no differencein the proliferation of cells in the WKM of WT and double-mutantfish at 4 mpf, as shown by a BrdU incorporation assay (Fig. 5C).We also examined the congenital phenotypes in the double

mutants. The sex reversal seen in rad51 single mutants was cor-rected (SI Appendix, Fig. S7 A–C), but the size defect was not (SIAppendix, Fig. S7D). However, neither female, nor male doublemutants were fertile (SI Appendix, Fig. S7 A–C). Taken together,our data suggest that the marrow hypocellularity in adult fish wasnot because of the smaller size of rad51mutant, meaning that thesetwo phenotypes are uncoupled. Instead, we observed a high cor-relation between the number of HSPCs generated early duringembryonic development and the kidney marrow cellularity inadulthood. Therefore, the rescue in the embryonic definitive he-matopoiesis can revert all defects observed in adult hematopoiesisin rad51 mutants.Finally, the tumor incidence of the double mutants was 30%,

with the first tumors developing from 5 mpf [5 mo earlier thanreported for p53 single mutants (43)]. The tumors resembled ma-lignant peripheral nerve sheath tumors (MPNSTs), (SI Appendix,Fig. S7 E and F), which is the most common type of malignancy inp53−/− fish (43). None of the other fish (i.e., WT, rad51 mutants,and heterozygotes) developed any kind of noticeable tumor, in-cluding the oldest 13-mpf rad51−/− fish.

rad51−/− Fish Are More Sensitive to Inflammatory Stress. An aber-rant inflammatory response has been postulated to be one of thepotential causes of the BMF in FA. This is thought to be a result ofincreased expression of inflammatory cytokines in FA patients andan excess apoptosis of HSPCs in response to these factors. To testthis hypothesis, we developed a zebrafish model of prolonged in-flammatory stress. Over a period of 4 wk, we injected rad51−/− andWT fish with the immunostimulant polyinosinic:polycytidylic acid(pI:pC) and assessed changes in the kidney cellularity, lineageoutput, and the expression of genes associated with inflammation(Fig. 6A).PI:pC resembles double-stranded RNA and is known to induce

the expression of proinflammatory cytokines, such as TNF-α (44)and IL-1 (45). Therefore, it has been considered to accuratelymimic viral infection (45) in murine and fish models (44–47). In-deed, a single intraperitoneal injection of pI:pC in zebrafish induced

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Fig. 3. HSPCs in the kidney adult rad51 mutant fish show increased pro-liferation. (A) AV-PI assay to assess apoptosis in the kidney. Two-sided t test,n+/+ = 4, n−/− = 4. (B) Schematic of the experimental design for the BrdUincorporation experiments. Fish were injected once with 10 mg/mL BrdUand culled after 1, 7, or 14 d to obtain the blood and kidney marrow forantibody staining and FACS analysis. (C) Percentage of BrdU+ cells in thekidney (i). Two-sided Student’s t test, P = 0.024 at 1 d postinfection (dpi)and P > 0.05 at 14 dpi; 1 dpi n+/+ = 5, n−/− = 6; 7 dpi n+/+ = 6, n−/− = 6; 14 dpin+/+ = 5, n−/− = 5. Percentage of BrdU+ cells in the peripheral blood (ii). Two-sided Student’s t test, P = 0.0015 at 1 dpi and P > 0.05 at 14 dpi; 1 dpi n+/+ = 5,n−/− = 6; 7 dpi n+/+ = 6, n−/− = 5; 14 dpi n+/+ = 5, n−/− =5. (D) Percentage of gata1:GFP+ cells in the kidney at 4 mpf. Two-sided Student’s t test, P = 0.025, n+/+ = 6,n−/−= 6. (E) Percentage of dim and bright cd41:GFP+ cells in the kidney at 4 mpf,labeling thrombocytic progenitors, and mature thrombocytes, respectively.Two-tailed Student’s t test. Thrombocytic progenitors: P = 0.023, maturethrombocytes: P = not significant, n+/+ = 10, n−/− = 10. Bars represent mean ±SEM in all graphs; n.s., not significant.

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robust inflammatory response just 6 h postinjection (SI Appendix,Fig. S8A). Mutants did not express more inflammatory cytokineswhen unchallenged (SI Appendix, Fig. S8B). Importantly, the pro-longed exposure of adult WT fish to pI:pC resulted in a twofoldincreased production of monocytes (Fig. 6B) but did not overtlyaffected kidney marrow cellularity or cell viability (Fig. 6 C and D).In addition, we observed clear up-regulation of monocyte-specificgenes (SI Appendix, Fig. S8C). The skew toward monocyte pro-duction in turn decreased the erythrocyte output and to a lesserextent other lineages. In contrast, repeated pI:pC injections ofrad51−/− fish led to no significant change in the number of producedmonocytes (Fig. 6B) and an ∼25% decrease in the total number ofcells in the kidney (Fig. 6C). As in the WT fish, the prolonged in-flammation decreased the number of erythrocytes in the kidneymarrow of rad51−/− fish, thus possibly contributing to the overallreduction in kidney cellularity. This was mediated by a normaliza-tion of proliferation rates in the kidneys of pI:pC-injected mutantfish (Fig. 6E and SI Appendix, Fig. S8D). Therefore, the prolongedinflammatory stress can lead to severe marrow defects in ourrad51−/− zebrafish FA model not only in terms of the lineage out-put, but also by affecting the total marrow cellularity.

Interestingly, the unchallenged rad51 mutants down-regulatedp53, but showed an exaggerated reaction in p53 expression to re-peated inflammatory stress (Fig. 6 F, i). Fitting with this observa-tion, p21 expression followed a similar trend in gene expression innoninjected and pI:pC-injected mutants (Fig. 6 F, ii).

rad51−/− WKM Cells Are Unaffected by Acetaldehyde-Induced Stress.DNA damage induced by small aldehydes has been proposed as amajor cause of exhaustion of the HSCs in the BM of FA patients(12–18). To examine the sensitivity of rad51 mutants to acetalde-hyde, we exposed them to acetaldehyde-induced stress over a pe-riod of 4 wk (SI Appendix, Fig. S9A). Our analysis revealed anincrease in WKM cellularity in WT fish in all blood cell types (SIAppendix, Fig. S9 B and C) upon acetaldehyde injections, whereasrad51 mutant cell numbers were unaffected. The viability of WKMcells was decreased to a similar extent in both WT and mutant fish(SI Appendix, Fig. S9D) but with no change in p53 expression in theWKM (SI Appendix, Fig. S9E).

DiscussionThe FA genes encode proteins that function cooperatively in theFanconi DNA-repair pathway. Here we characterized a viable

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Fig. 4. The rad51sa23805 HSPC defect starts during embryonic development. (A) ISH using a cmyb-specific probe at 2 dpf; the arrow shows HSPCs. Representativeimages of the three different staining categories are shown (i) and a quantification of the different genotypes (ii) n = 119 from two clutches. (B) ISH using a cmyb-specific probe at 4 dpf; the arrow shows HSPCs. Representative images of the three different staining categories are shown (i) and a quantification of the differentgenotypes (ii), n = 120 from two clutches. (C) Quantification of BrdU+ cells in the tail at 2 dpf. Two-sided Student’s t test, P = 0.042, n+/+ = 3, n−/− = 3.(D) Quantification of BrdU+ cells in the CHT at 4 dpf. Two-sided Student’s t test, n+/+ = 4, n−/− = 4. Bars represent mean ± SEM in C and D. n.s., not significant. (E)Representative images of TUNEL-stained 2 dpf embryos from a rad51+/− in-cross. Dotted lines indicate the area of the CHT that was scored. Arrows indicateTUNEL+ cells. (F) Quantification of three clutches of TUNEL-stained 2 dpf rad51+/− in-crosses. Each clutch was scored blindly and consisted of 10+/+ and 10−/−

embryos each. Shown is the mean of all clutches ± SEM. (Magnification, 100× in all images.)

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vertebrate rad51 loss-of-function mutant. Loss of rad51 in zebrafishrecapitulated many congenital features of FA, such as short statureand microphthalmia (1), as well as hematological defects, includingmarrow cytopenia and accumulation of macrocytic erythrocytes incirculation (1, 7). Most importantly, rad51−/− fish showed increasedsensitivity to cross-linking agents, an absolute diagnostic criterionof FA (7).Furthermore, our results show that loss of rad51 does not lead to

higher sensitivity to PARP inhibitors. Previous research on mam-malian cells showed that BRCA1/2 mutant cells are more sensitiveto PARP inhibition. One model proposes that this is because of theimportance of PARP-1 in reactivating stalled replication forks inHR-deficient cells (48). That model is not compatible with ourdata, suggesting that PARP-BRCA synthetic lethality stems fromfunctions unrelated to HR. PARP1 is highly conserved betweenzebrafish and humans (∼70%). A small-molecule inhibitor such asDiQ should therefore be able to enter the zebrafish embryos andinhibit the enzyme the same way as in human cells. Further evi-dence for this is provided by the lethality of high doses of this drugin WT and rad51 mutant fish. However, we were unable to obtainbrca2 mutant zebrafish to use as a positive control for DNAdamage in response to PARP inhibition. This should be taken intoaccount when interpreting our findings.

Although loss of Rad51 leads to an early embryonic death inmice (33, 34), zebrafish lacking rad51 survive to adulthood. This isnot entirely surprising, as fish lacking brca2 also survive to adult-hood (49, 50), whereas Brca2mutant mice die before birth (51, 52).It has been hypothesized that maternal mRNA contributes toembryonic viability of zebrafish mutants (49). This seems likely,considering the presence of maternally derived Rad51 in rad51mutant embryos. However, another plausible explanation is thefunctional redundancy between rad51 and its paralog rad51l1 inzebrafish. Our analysis suggests that rad51 and rad51l1 are able topartially compensate for each other, leading to the lethality ofdouble mutants. In contrast, NHEJ did not appear to play a role incompensating for the loss of HR. Interestingly, rad51−/− fish, likebrca2−/− fish, show sex reversal that can be rescued upon p53comutation (49, 50). Additional p53 loss further caused develop-ment of MPNSTs in both brca2 and rad51mutants, which occurredconsiderably earlier and at the higher incidence than in p53 singlemutants (43, 49, 50, 53).FA is genetically and phenotypically heterogeneous disorder, but

BM failure is the most common cause of death (1), with patientshaving lowered CD34+ progenitor cell numbers from birth (10, 54).This led to the hypothesis that hematological defects in FA origi-nate from an impairment of HSPCs during embryonic develop-ment, which leads to a decreased number of HSPCs at birth (10,11). Here we provide in vivo evidence that the decrease in HSPCnumbers in adult fish indeed stems from a combination of de-creased proliferation and increased apoptosis during embryonicdevelopment. This defect appears to be mediated via p53 (10), asour p53/rad51 double mutants did not display any observable he-matological defects in embryos or adults.In agreement with our study, knockdown of fancd2 in zebrafish

embryos causes massive apoptosis in the whole body, associatedwith up-regulation of genes in the p53 pathway, as well as de-creased expression of cyclins. Strikingly, co-knockdown of p53rescued both the cyclin down-regulation, as well as apoptosis (55),resembling the situation in our fish and providing further evidencefor the importance of p53 signaling. This phenotype is, however,only partly consistent with what was observed in murine fetalFancd2−/− cells. In the murine model the reduced number ofHSPCs was set off by p38-mediated reduced proliferation, withoutany involvement of apoptosis or p53 signaling (56). More researchinto the causes of the early HSPC defects and how they can bemitigated is warranted. Because of the external development andtheir transparency, zebrafish embryos would be an ideal tool todiscover compounds that can alleviate these defects.The decreased WKM cellularity in rad51 zebrafish mutants also

mirrors defects seen in Lig4 (a protein involved in NHEJ) mutantmice, which show decreased BM cell numbers, coupled to an ap-proximately twofold increase in proliferation of long-term HSCs(57). This underlines the importance of repairing double-strandedbreaks in HSC maintenance and suggests hyperproliferation ofblood progenitors might be a common mechanism to cope withdecreased cell numbers in the kidney/BM. However, like mostmurine FA models (58, 59), the fish never progressed to pancy-topenia or spontaneous BMF/kidney marrow failure, which isprobably because of species differences in the HSC compartment,lifespan, and rearing conditions. Nevertheless, together with theclinical data (29, 30), these facts provide further evidence for thedesignation of RAD51 as FANCR and show that our rad51−/− fishare a suitable model for FA.The progressive decline of HSC numbers in FA patients leads to

BMF and there is considerable evidence for an overproduction ofinflammatory cytokines in patients and mouse models (26, 60–64).Furthermore, inflammatory stress (for example by repeated pI:pCinjections) can induce BMF in mouse models of FA (65, 66). Wedid not observe an increase in inflammatory cytokines in our un-challenged rad51 mutants, ruling out cytokine overproduction as acause for hematological defects. We did, however, observe a 25%

CB

A

p53+/+ rad51+/+

p53-/- rad51+/+

p53-/- rad51-/-

cmyb - 4 dpf

p53+/+ rad51-/-

Fig. 5. The HSPC defects in rad51sa23805 fish are rescued in a p53 mutantbackground. (A) Representative images of 4-dpf embryos resulting from in-crosses of p53+/− rad51+/− parents stained using a cmyb-specific probe. Thetotal number of embryos used (all genotypes) n = 237 from four clutches.For information about all genotypes, see SI Appendix, Table S3. Arrowsindicate HSPCs. (Magnification, 100×.) (B) Percentage of BrdU+ cells in thekidney at 4 mpf at 1 dpi. Two-sided Student’s t test, np53

+/+rad51

+/+ = 5,np53

−/−rad51

−/− = 5. (C ) Number of total cells per kidney at 4 mpf quantifiedusing a hemocytometer. Analysis using one-way ANOVA [F(3, 43) = 10.45,P < 0.0001], individual P values shown in the figure are from Tukey’s post hoctest, np53

+/+rad51

+/+ = 16, np53+/+

rad51−/− = 16, np53

−/−rad51

+/+ = 6, np53−/−

rad51−/− = 8. Bars represent mean ± SEM in B and C; n.s., not significant.

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decrease in WKM cellularity after repeated of pI:pC injections,possibly because of decreased proliferation compared with un-challenged fish. Unlike WT fish, rad51 mutants were unable torespond appropriately to inflammation and increase monocyteproduction. The decrease in WKM cellularity as a result of in-flammatory stress was accompanied by an up-regulation of p53,again underlining the high importance of this pathway in the eti-ology of FA and explaining the normalized proliferation. Interest-ingly, we were unable to elicit similar changes using acetaldehyde,indicating that damage induced by small aldehydes is not a majorfactor in FA pathogenesis, at least after birth.Our study characterized a viable vertebrate model of RAD51

loss, which recapitulates many human FA symptoms and is thusalso a zebrafish model of the disease. Further study of this mutantwill increase our knowledge of the cellular roles of RAD51 in vivoand will deepen our understanding of the molecular pathology ofFA. Transparency of zebrafish embryos, their high fecundity, andthe existence of transgenic lines labeling various blood lineagesmakes them very amenable for high-throughput screening incomparison with other model organisms. The application of therad51 mutant line will significantly impact the development ofnovel therapeutics to improve HSPC function in FA patients, by

screening for molecules that can alleviate the HSPC reductionin embryos.

MethodsZebrafish Care and Strains. Fish lines were maintained in the Sanger Institutezebrafish facility according to European Union regulations. WT fish wereof the Tübingen long-fin strain. Fish were genotyped as described pre-viously (35).

Western Blotting. Western blotting was carried out as described previously(67). Antibodies can be found in SI Appendix, Table S1.

Embryo Irradiation. Embryos were irradiated at 24 hpf in a Gammacell 1000Elite Blood Irradiator (MDS Nordiron) at 750 cGy.

Immunostaining. Staining was carried out as described previously (68). We usedHoechst 33342 as nuclear stain. Embryos were imaged on a Leica SP-5 confocalmicroscope using a 40× water-immersion lens. Antibodies can be found in SIAppendix, Table S1.

Chromosome Spreads. Embryos were treated with 1 μg/mL DEB (SigmaAldrich), 5 μg/mL MMC (Sigma Aldrich), 1 nM CPT (Sigma Aldrich), or 10 μM1,5-isoquinolinediol (Sigma Aldrich) in egg water between 4 and 24 hpf.From here on we followed The Zebrafish Book, 4th edition (69), but kept

rad51+/+ or -/-

pI:pC via IP

0 7 14 21 24

Cell CountsFACSqPCRs

Cull

A B C

E i iiFD

Fig. 6. Lack of Rad51 causes increased sensitivity to prolonged inflammatory stress. (A) Schematic of the experimental design. Both WT and rad51−/− fishwere injected every 7 d with 10 μL 10 mg/mL pI:pC acid, four injections in total. All fish were culled 3 d after the last injection. Control fish were not injected;Rad51+/+, nnoninjected = 10, ninjected = 9; Rad51−/−, nnoninjected = 8, ninjected = 9. (B) Absolute number of cells belonging to different blood lineages in the kidneygained by combining FACS data with the cell counts shown in A. Statistical tests were carried out individually for each cell type, using two-way ANOVA.P value shown on the graph stems from a post hoc �Sidak multiple-comparison test, comparing noninjected to injected fish within each genotype. For allgroups, n is the same as in A. (C) The total number of cells in the kidney in injected and noninjected fish. Two-way ANOVA was carried out on the reciprocal ofthe data to fulfill the requirement of homoscedasticity as measured by Bartlett’s test (before transformation: P = 0.0002, after transformation: P = 0.095).There was a statistically significant effect of mutation status [F(1, 32) = 29.86, P < 0.0001] and of injection status [F(1, 32) = 6.778, P = 0.014]. P value shown onthe graph stems from a post hoc Tukey multiple-comparison test. For all groups, n is the same as in A. (D) Viable cells as determined by PI-staining. Two-wayANOVA revealed a significant influence of injection status [F(1, 32) = 100.1, P < 0.0001]. P values on the graph stem from a post hoc Tukey multiple-comparison test. Bars represent mean ± SEM in B–D. (E) Percentage of BrdU+ cells in the WKM of WT and mutant (KO) fish in response to pI:pC. 1I, oneinjection; 2I, two injections; 4I, four injections. (F) Relative expression of genes linked to apoptosis and proliferation. (i) p53. P value shown on the graph stemsfrom a post hoc Tukey multiple-comparison test. (ii) p21. Bars represent geometric mean ± 95% CI in E and F.

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the head of each embryo for genotyping. VECTASHIELD mounting me-dium with DAPI was used to visualize the spreads.

NHEJ Inhibition. Embryos were treated with 1, 10, 25, 50, 75, and 100 μMSCR-7(Sigma Aldrich) at 4 hpf. Embryos were scored for defects at 24 hpf.

CRISPR-Cas9. Mutations in rad51l1 were induced at exon two, leading to a 7-bpdeletion (478-484delTGGGTCC in the cDNA). The targeted DNA sequence was5′-GGATGTCCTGTCGGTCACCCAGG-3′. ssDNA oligonucleotides 5′-TAGGATGTCC-TGTCGGTCACCC-3′ and 5′-AAACGGGTGACCGACAGGACAT-3′ (Sigma-Aldrich)were annealed and ligated with pDR274 vector (Addgene) linearized with BsaI(New England Biolabs) to make guide RNA (gRNA) expression vectors. gRNAwasprepared with MAXIscript T7 kit (Life Technologies) using DraI-linearized gRNAexpression vector as a template, and Cas9 mRNA was synthesized usingmMESSAGE mMACHINE T7 kit (Ambion) and pMLM3613 expression vector(Addgene) linearized with PmeI (New England Biolabs). Zebrafish embryos wereinjected at the one-cell stage with 12.5 pg of gRNA and 160 pg of Cas9 mRNA.For a detailed zebrafish CRISPR methodology, see Brocal et al. (70).

Histology. Formalin-fixed tissues were processed and sectioned using stan-dard techniques (71), followed by staining with Harris H&E.

AV-PI Assay. We used the Alexa Fluor-488 Annexin V/Dead Cell Apoptosis Kit(Thermo Fisher) according to the manufacturer’s instructions.

BrdU Assay on Adults. Fish were injected with 10 μL of 10 mg/mL BrdU (SigmaAldrich) and culled at 1, 7, or 14 d postinjection. We extracted kidney andblood, made single-cell suspensions, and fixed cells in 70% EtOH overnight.BrdU immunostaining was carried out as described previously (67), after whichthe cells were resuspended in PBS and analyzed using FACS.

BrdU Assay on Embryos. Embryos were chilled on ice for 15 min. This was fol-lowed by a 20-min incubation in 10 mMBrdU on ice. After a 3-h recovery in eggwater, embryos were fixed in 4% PFA. Heads from embryos were used forgenotyping. For 2-dpf embryos, the whole tails were pooled according to ge-notype, whereas for 4-dpf embryos just the CHT was dissected. Samples weretreated with 10 mM DTT in 1× Danieau’s solution for 30 min at room tem-perature, followed by incubation in liberase (Roche) in PBS for 3 h at 37 °C. Thereaction was stopped by replacing the solution with 5% FBS/PBS. Single-cellsuspensions were fixed in 70% ethanol overnight. From here on, the staining

process and analysis was identical to cells obtained from adults. Antibodies canbe found in SI Appendix, Table S1.

TUNEL Assays. Embryos for TUNEL assays were fixed and stored as for ISH.Staining was carried out using the In-Situ Cell Death Detection Kit, AP (Roche),according to the manufacturer’s instructions.

Kidney FACS. Dissected kidneys were placed in 5% FBS/PBS and processed tosingle-cell suspensions. Dead cells were excluded using DAPI (Sigma-Aldrich) orPI. Flow cytometry was carried out on a MoFlo XDP (Beckman Coulter), a BDLSRFortessa, or a BD Influx (BD Biosciences).

In Situ Hybridization. ISHwas carried out as described previously (72). We used acmyb antisense probe. Embryos were blindly sorted into high-, medium-, andlow-staining conditions followed by genotyping.

Long-Term pI:pC Injections. Fish were injected with 10 mg/mL pI:pC (Sigma-Aldrich) once a week, totaling four injections. Fish were culled 1 or 3 d afterthe last injection. Optionally, 10 mg/mL BrdU were added to the last injection.Kidneys were processed to single-cell suspensions for FACS, as described above,and cell numbers counted. Remaining cell suspensions were used for gene-expression analysis.

Long-Term Acetaldehyde Injections. For long-term acetaldehyde injections, thesame protocol as for the pI:pC injections was followed, but using 10 μL 1%acetaldehyde (Sigma Aldrich) in PBS instead for the injections.

qPCRs. The qPCR reaction used SYBR green (Thermo Fisher) and run on aQuantStudio 3 (Thermo Fisher) qPCRmachine. Statisticswere carriedout on rawΔCt values. Primers used are listed in SI Appendix, Table S2.

ACKNOWLEDGMENTS. We thank the Sanger Institute Zebrafish MutationProject for supplying the rad51sa23805 allele; Sebastian Gerety for supplyingthe tp53zdf1 line; Yvette Hooks for her help with histology; and the SangerInstitute FACS core facility and Charlotte Labalette for their experimental help.This work was supported by Cancer Research UK Grant C45041/A14953 (toA.C.); a core support grant from the Wellcome Trust and Medical ResearchCouncil to the Wellcome Trust–Medical Research Council Cambridge Stem CellInstitute; and a European Hematology Association–Jose Carreras FoundationYoung Investigator Award and Isaac Newton Trust grant (to A.C.).

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