A model for reticular dysgenesis shows impaired sensory organ development and hair cell regeneration linked to cellular stress

Mutations in the gene AK2 are responsible for Reticular Dysgenesis (RD), a rare and severe form of primary immunodeficiency in children. RD patients have a severely shortened life expectancy and without treatment die a few weeks after birth. The only available therapeutic option for RD is bone marrow transplantation. To gain insight into the pathophysiology of RD, we previously created zebrafish models for an AK2 deficiency. One of the clinical features of RD is hearing loss, but its pathology and causes have not been determined. In adult mammals, sensory hair cells of the inner ear do not regenerate; however, their regeneration has been observed in several non-mammalian vertebrates, including zebrafish. Therefore, we use our RD zebrafish models to determine if AK2 deficiency affects sensory organ development and/or hair cell regeneration. Our studies indicated that AK2 is required for the correct development, survival and regeneration of sensory hair cells. Interestingly, AK2 deficiency induces the expression of several oxidative stress markers and it triggers an increased level of cell death in the hair cells. Finally, we show that glutathione treatment can partially rescue hair cell development in the sensory organs in our RD models, pointing to the potential use of antioxidants as a supportive therapeutic modality for RD patients, not only to increase their chances of survival, but to prevent or ameliorate their sensorineural hearing deficits.


Introduction
Reticular dysgenesis (RD) is a rare form of severe combined immunodeficiencies (SCID), a heterogenous group of immunological diseases usually characterized by profound defects to the T and B lymphoid lineages(1, 2). In particular, RD patients also present a reduced number of neutrophils, with a specific block at the promyelocyte stage, and an unresponsiveness to GCSF stimulation in combination with severe sensorineural hearing loss (3,4). The immunodeficiencies associated with strong neutropenia leads these patients to recurrent severe infections and premature death within the first weeks or months of life. For these patients, a hematopoietic stem cell transplantation (HSCT) represents the only available treatment, but, because of the severity of the disease, pre-conditioning is necessary in order to increase the chances of engraftment of the donor stem cells (5). However, the HSCT does not improve the non-hematological defects such as the hearing loss, which has a huge impact on the cognitive functioning and quality of life of these children (6,7).
Mutations in the adenylate kinase 2 (AK2) gene have been shown to be responsible for the disease(1, 2,8). AK2 is an enzyme localized to the intermembrane space of the mitochondria, where it plays an important role in sustaining mitochondrial respiration and cellular energy metabolism (9)(10)(11). Because the knockout of mouse AK2 showed an early embryonic lethality (8,12), other cellular and animal models needed to be developed. Insect models of AK2 deficiency indicated an essential role of the gene in embryonic growth and cell survival (13,14); moreover, they suggested that maternal AK2 mRNA can, at least initially, compensate for the lack of AK2 gene zygotic transcription. In zebrafish, AK2 knock-down induced by MO injection showed hematopoietic defects without affecting general embryonic development (2,8). These results were confirmed by two different mutant lines carrying a frameshift mutation in zebrafish AK2 exon 1 and a missense mutation in exon 4 (8). Similar to what was observed in vitro in patient fibroblasts and CD34+ bone marrow cells (2,11), zebrafish mutants presented an increased level of cellular oxidative stress leading to apoptosis and cell death (8). Notably, these phenotypes can be reduced by the administration of antioxidants in both zebrafish mutants and, moreover, the same kind of treatment was able to rescue myeloid differentiation in vitro in induced pluripotent stem cells (iPSCs) obtained from fibroblasts of a RD patient (8).
Although most of the work linked the AK2 role to its bio-energetic activity, other evidence highlighted the presence of alternative functions, partially unrelated to its enzymatic activity and/or the mitochondrial localization (5). AK2 protein has been found to associate with dual-specificity phosphatase 26 (DUSP26), resulting in the suppression of cell proliferation by FADD dephosphorylation (12). In addition, it has been shown to be involved in an amplification loop that ensures the execution of the intrinsic apoptosis via an interaction with FADD and caspase 10 (15). Previous reports showed that an AK2 deficiency impairs the regular induction of the unfolded protein response (UPR) mechanism in vitro (16,17). Finally, using RD patient-derived iPSCs, recent work showed a reduction of nuclear ATP levels in AK2-deficient cells during specific stages of hematopoietic differentiation (10). Reduced levels of nuclear ATP could be responsible for the altered transcriptional profile observed during hematopoietic differentiation (10,11). Overall all these studies collectively seem to suggest that, at least to some extent, the cellular AK2 roles can be cell-type or context specific.
Sensorineural hearing loss is the most common form of human hearing loss and it can be due to several different factors including genetic mutations, ototoxic compound exposure, aging, infectious diseases, or environmental stress, such as prolonged exposure to excessive noise (6,18). In general, all these different factors can induce damage to the mechanosensory hair cells in the organ of Corti or the stria vascularis and they can also impair the function of the spiral ganglion neurons or of the more proximal auditory structures (19). Because of the limited regenerative ability of mammals, hair cells cannot regenerate after damage and the resultant hearing loss is permanent. In contrast, non-mammalian vertebrates like zebrafish possess a huge regenerative potential and they can replenish hair cells during homeostasis or after damage, providing a model to study hair cells development and, in particular, hearing restoration after damage (20)(21)(22).
Although zebrafish inner ears lack a structure strictly equivalent to the mammalian cochlea, most of the functions present in all other vertebrates (i.e. hearing and vestibular functions) are fully conserved (22). Moreover, zebrafish possess another 5 mechanosensory system on their body surface called the lateral line, which provides feedback information about water flow along the body of the fish (23)(24)(25). The lateral line is composed of a network of several small sense organs, the neuromasts, distributed over the zebrafish body in a specific pattern (23). It comprises two major branches, an anterior part that extends on the head (anterior lateral line or ALL), and a posterior part on the trunk and tail (PLL). Each neuromast consists of a central core of hair cells which are innervated by lateralis afferent neurons. The hair cells are then surrounded by two different types of non-sensory cells: supporting and mantle cells (23). Because of its structural simplicity and experimental accessibility, the lateral line, and in particular the PLL, has become a very popular model to study hair cells development and regeneration and it contributed significantly to the understanding of the molecular pathways that control those phenomena (26).
A previous study using adenylate kinase 2 immunostaining on mouse inner ears, indicated that AK2 protein localization was specifically limited to the lumen of the capillaries of the stria vascularis only in 7 days post-natal mouse embryos(1). Notably, no signal has been observed in other blood vessels or other cochlear structures such as the Organ of Corti(1). The stria vascularis is the portion of the inner ear responsible for the endolymph production, which is important for the metabolic support of the Organ of Corti(1,19). Therefore, it has been postulated that, in those specific structures, AK2 could work as an ecto-enzyme, controlling the ATP and/or potassium concentrations in the endolymph(1). As a consequence, its deficiency could result in hearing loss through a failure in the regulation of the potassium-and/ or ATP-homeostasis of the endolymph, as observed in the case of mouse connexin30 null mutants (27).
To shed light on the causes of RD hearing loss, we used the zebrafish models of RD (8).
Our data showed that, zebrafish ak2 is expressed in inner ear and neuromasts structures and its deficiency severely affects the survival of hair cells in those structures through an increased level of oxidative stress and cell death. Finally, the administration of glutathione (GSH) to zebrafish ak2 mutants showed that antioxidant treatments were able to partially increase the number of hair cells in inner ear and PLL of ak2-deficient animals and this increase was both at the level of development and long-term survival of the hair cells.

The Zebrafish adenylate kinase 2 gene is expressed in sensory organs
Reticular dysgenesis is a rare hematological disorder caused by mutations in the adenylate kinase 2 (AK2) gene(1, 2). Although at least one RD patient with skeletal defects has been reported so far (28), the only non-hematological clinical feature required for the diagnosis of RD is the sensorineural deafness/hearing disability (4,5).
Because of the paucity of data, overall the pathophysiology of the hearing loss in RD patients is essentially uncharacterized. Here we used our zebrafish models of RD to better understand the physiopathology of hearing loss in RD patients. Initially we tested if the zebrafish AK2 gene (ak2) was expressed in sensory organs such as the inner ear or the lateral line system. Whole mount in situ hybridization (WISH) analysis of endogenous ak2 expression during zebrafish development showed that from 3 days post fertilization (dpf) ak2 gene is expressed in different anatomical regions, included the otic vesicle ( Figure 1A). To better characterize the territories of ak2 expression in the otic vesicle, we performed transversal sectioning on 5 dpf stained embryos. As shown in Figure 1 These data suggest that, in zebrafish, ak2 is expressed in sensory organs and that the lack of its expression could potentially affect inner ear or neuromast development to some extent.

Lack of ak2 affects inner ear hair cell development
In a previous study, we showed that two different ak2 mutants, carrying a missense mutation (previously denoted as ak2 L124P/L124P , and here indicated as ak2 hg16 ) or frame- 7 shift mutations in exon1 of ak2 (ak2 del2/del2 and ak2 ins4/ins4 mutants, here designated as ak2 hg14 and ak2 hg15 , respectively), presented very similar severe hematopoietic phenotypes affecting hematopoietic stem and precursor cell development (8). However, survival analysis showed that some ak2 hg16 mutants can reach adulthood, although with a lower than expected ratio of survivors (~10% compared to the expected percentage of ~25%) ( Figure 1B, left panel). In contrast, ak2-deficient mutants ak2 hg14 and ak2 hg15 , lacking the ak2 mRNAs(8), start to die around 5 dpf; and by 11 dpf we were not able to find any surviving homozygous mutants ( Figure 1B, right panel and data not shown).
These data suggest that, despite the severity of the hematopoietic defects observed in all ak2 mutants, ak2 hg16 likely represents a hypomorphic mutation, while the other two alleles (ak2 hg14 and ak2 hg15 ) are more likely fully null. Notably, in ak2 hg16 adults we did not observe any behavioral signs indicating potential sensorineural defects (22,30) such as "circling" or the lack of a startle response. When we compared hair cell bundle density of inner ear structures from ak2 hg16 and their heterozygous siblings (ak2 hg16/+ ) of the same age, we did not find a statistically significant difference ( Figure 1C and Figure   1-figure supplement 2). However, using the Tg(pou4f3:GAP-GFP) transgenic line to visualize the mature hair cells, we found a statistically significant difference in the average number of hair cells in 3 and 4 dpf developing inner ears of ak2 hg14 mutants compared to their control siblings ( Figure 1D).
Taken together, these data suggest that a more severe ak2 deficiency is needed to impair the correct development of the hair cells in the zebrafish inner ears compared to the hematopoietic defects seen in the hypomorphic allele.

ak2 deficiency affects posterior lateral line development
One hypothesis for the sensorineural hearing loss in AK2-deficient patients is that the composition of the luminal space is compromised, resulting in hair cell loss(1, 5). To understand if hair cells in a different context (i.e. not in a luminal space) could similarly be affected by the lack of ak2 expression, we examined the posterior lateral line (PLL) of our knockout fish. In zebrafish larvae, the PLL possess at least three different populations of neuromasts denoted as primary, secondary and intercalary (IC) (24). Primary neuromasts are deposited by the first migrating primordium (primI) which travels from the otic vesicle along the midline of the trunk on both sides of the embryo (23,31,32) to the tail, regularly depositing primordial neuromasts. Moreover, all the primary neuromasts are connected by a thin continuous stripe of cells (interneuromast cells or INCs) which, later during development, act as multipotent stem cells to produce intercalary neuromasts (33). By 2 dpf, the migration of the first primordium is complete as it reaches the most caudal part of the trunk of the embryos.
At approximately the same time, a second, slower primordium (primII) starts its migration from the otic vesicle following a similar route used by the primI and it generates a second set of neuromasts called "secondary neuromasts" (or LII) (24). In ak2 hg16 hypomorphic embryos we did not observe any major defects to the PLL formation as indicated by alkaline phosphatase (AP) staining(34) at 4-5 dpf or Yo-PRO-1 staining (35,36) of functional hair cells from 3 to 5 dpf ( function has a negative impact on the regeneration of hair cells, we checked if ak2 expression was present after hair cells ablation with copper sulfate. WISH analysis performed at different time points after complete hair cell ablation with a 1-hour treatment with 10 µM copper sulfate, showed that ak2 expression was increased in neuromasts 3 hours post ablation (hpa) (data not shown). Accordingly, we also observed a strong reduction of Yo-PRO-1 + hair cells in ak2 hg14 embryos at 2 days post ablation (dpa) compared to control siblings ( Figure 2D). Moreover, we observed similar results performing copper sulfate hair cell ablation assay on ak2 hg14 in Tg However, from ~4 dpf eya1 expression was undetectable in the secondary primordium ( Figure 3C) and strongly reduced or completely absent in LII.1 and LII.2 neuromasts ( Figure 3C).
For a more detailed study of the secondary neuromast formation and development in ak2-deficient embryos, we crossed ak2 hg14/+ and ak2 hg15/+ animals with transgenic lines marking different cellular components of neuromasts. As previously observed using Yo-PRO-1 staining ( Figure 2B), the analysis of ak2 hg14 Tg(pou4f3:GAP-GFP) embryos further confirmed the lack of GFP + cells (corresponding to hair cells) in secondary neuromasts from ~4 dpf ( Figure 3D). Notably, at 3dpf the confocal analysis showed the presence of GFP + cells in LII.1 neuromasts, although with a severely reduced number of cells compared to the controls ( Figure 3D, yellow arrows). The lack of Yo-PRO-1 positive cells in LII.1 of ak2 homozygous mutants at the same stage ( Figure 2B) suggested these GFP positive hair cells are not fully mature and functional, despite expressing pou4f3.
Using the double transgenic line Tg(tnks1bp1:EGFP; atoh1a:dTOM)(39) to label immature hair cells (red) and supporting cells (green), we were able to highlight further defects in secondary neuromast formation ( Figure 3D). In particular, the analysis showed that ak2-deficient embryos presented a specific lack of both the dTOM and EGFP signals in LII.1 neuromasts from ~3 dpf on. Although the vast majority of the embryos analyzed did not present any positive signal in the secondary neuromasts at 5 dpf, surprisingly, in sporadic animals we observed dTOM + cells between L1 and L2 neuromasts ( Figure 3D, white arrowhead). Notably, at the same stage we never observed more mature hair cells (marked by pou4f3:GAP-GFP + ) and, more importantly, these rare atoh1a:dTOM + cells were never surrounded by tnks1bp1:EGFP + supporting cells suggesting a depletion of the precursor cell pool used to populate the secondary neuromasts. This is similar to the depletion of hematopoietic stem and precursors cell observed in the zebrafish ak2 alleles (8).
These data indicate that ak2-deficient embryos present an altered expression of markers for different neuromast cell populations, with a reduction in both immature and mature hair cell-and supporting cell-specific markers. However, the sporadic presence of solitary atoh1a:dTOM + cells ( Figure 3D, white arrow), in addition to the persistence in WISH analysis of a faint signal for the eya1 marker in secondary neuromasts at 5 dpf dpf showed that at 3 dpf some of the ak2-deficient embryos completely lacked EGFP signal in the LII.1 secondary neuromasts ( Figure 4A). However, in the remaining ak2 hg15 population where the LII.1 retains some EGFP positive signal, secondary neuromast development was severely affected. In particular, we never observed the formation of EGFP + LII.2 neuromasts. In addition, while wild-type LII.1 neuromasts migrate ventrally a few cell thicknesses, LII.1 neuromasts in ak2 deficient larvae failed to migrate. In contrast to wild-type or carrier siblings, the neuromasts remained close to the horizontal myoseptum, at the same level of primary neuromasts ( Figure 4A). Similar migratory defects in secondary neuromasts have only been previously described in the strauss mutants(41). Notably, at 5 dpf there is still a mixture of phenotypes among the ak2 hg15 embryos some that are totally negative for EGFP + LII.1 neuromasts and some that possess the first secondary neuromast but no more. To confirm that the observed phenotypes were not due to early defects of the secondary neuromasts or the inability of primII to migrate, we also performed confocal analysis on ak2 hg15 Tg(-8.0cldnb:LY-EGFP) mutants and their siblings from 3.5 to 4 dpf (Video 1 and 2, respectively and Figure 4B). As summarized in Figure 4B, we did not observe major differences during primII migration between ~3 and 4 dpf.
As previously mentioned, intercalary neuromasts derive from the proliferation and differentiation of the INCs, a population of precursor cells connecting the neuromasts of the lateral line that was deposited by the primI during its migration(43). In physiological conditions, this population of cells remains quiescent because of the presence of inhibitory signals deriving from glial cells(44, 45), and isolated IC neuromasts can only rarely be observed between L1 and L2 primary neuromasts in WT embryos (24,42).
However, it has been shown that, the lack of inhibitory signals induced by loss of glial cells and/or the PLL nerve(44, 45) can prompt an ectopic production of IC neuromasts(42) in the region. Moreover, a previous study also showed that the ablation of the secondary primordium primII induces the ectopic production of IC neuromasts in the same region (24). Notably, in ak2-deficient embryos we never observed sporadic IC neuromasts nor an increased number of IC neuromasts between L1 and L2 neuromasts Notably, from 4 dpf we detected an increased number of interneuromast cells between the L1 and L2 neuromasts (yellow asterisks), suggesting some active proliferation of the interneuromast cells derived from primI. In the ak2 hg14 mutants, primI-and primII-derived interneuromast cells and the primII primordium (yellow and white arrowheads and white asterisk, respectively) were mRFP1 + until 4 dpf. Moreover, only some of the null embryos had LII.1 neuromasts positive for hair and supporting cell markers, and it was at a very low level of expression ( Figure 4C). Finally, at 5 dpf ak2-deficient embryos lacked the expression of both GFP and mRFP1 reporters between L1 and L2 primary neuromasts ( Figure 4C) suggesting the cells are not viable long term.
Overall, these data suggest a possible defect in the activation of the interneuromast cells with a resulting deficiency in the formation of IC neuromasts. The mRFP1 expression in PLL secondary structures confirmed the presence of LII.1 secondary neuromasts in some of ak2-deficient embryos at least until 4 dpf but not after. These data are consistent with a premature depletion of the stem cell pool for the lateral line organs. 13

Partial impairment of Wnt/β-catenin and Fgf signaling in ak2-deficient mutants
To verify that in ak2-deficient mutants the Schwann cells of PLL did not possess any severe deficiencies, we tested the effects of pharmacological inhibition of ErbB signaling using the tyrosine kinase inhibitor AG1478 (49)  Because at the tested dose we always observed a uniform response in the control population, the partial penetrance observed in the null mutants seems to reflect a real heterogeneity of the ak2 hg14 population's response to the treatment and not just variation in inhibition levels. It is important to note that the induced neuromasts (white asterisks) were atoh1a:dTOM and tnks1bp1:EGFP double positive, although their size was strongly reduced compared to control siblings. Because we never observed the expression of these two markers in the secondary neuromasts of ak2 mutants, their presence in AG1478-induced mutant neuromasts suggests a difference in the potency of secondary and IC neuromast precursors, probably due to their separate cellular origins.
Although little is known about the signaling pathways that orchestrate IC neuromast formation, a previous study showed that it requires both Wnt/β-catenin and Fgf signaling pathways(42). In particular, while the Wnt/β-catenin signaling is required for initial interneuromast proliferation, Fgf signaling is crucial for subsequent organization of interneuromast cells into a rosette structure and for triggering cellular differentiation(42). To determine if these pathways were affected in ak2 mutants, we treated the embryos from an ak2 hg14/+ incross with 6-BIO, a potent inhibitor of the GSK-3 enzyme ( Figure 5B). To obtain a quantitative analysis, we performed an AP staining of the embryos and we included an AG1478 treatment as further internal control. 6-BIO has been shown to result in the strong activation of the Wnt/β-catenin pathway(50) and the formation of extra IC neuromasts in the PLL of treated zebrafish embryos(42). As shown in Figure 5B, in ak2 hg14 untreated or DMSO-treated embryos secondary or IC neuromasts between L1 and L2 were always AP negative. At the same time, 100% of the AG1478 and 6-BIO-treated control embryos developed extra IC neuromasts. It's important to note that, in order to avoid 6-BIO toxicity from 24 hours of administration, we used it at a concentration of 0.2 µM (which is 10 times lower than the previously published dose(42, 51)). After 3 days of treatment with the selected dose, we observed a strong induction of ectopic neuromasts in the trunk regions of control embryos, without any major morphological defects or reduced survival of the embryos. These results were very similar to those we obtained on control embryos with a 2 µM AG1418 treatment ( Figure 5B). As previously observed ( Figure 5A), the ak2-deficient embryos treated with the AG1478 compound could be classified in three different groups. The AG1478 administration induced the formation of ectopic intercalary neuromasts in ~42% (5/12), a partial response in ~33% (4/12), and a lack of intercalary neuromast formation in the remaining ak2-deficient population (25%, 3/12). Similarly, we found a partial penetrance using the GSK-3 inhibitor 6-BIO, as well. However, in this case we observed that half of the ak2-deficient population (12/24) was totally unresponsive to the treatment, while the other half showed a very mild response, with a reduced number of ectopic IC neuromasts compare to controls. In both the treatments we noticed that, again, the ectopic IC neuromasts were significantly reduced in their size ( Figure 5A-B).
Overall these data suggest an impairment of the Wnt/β-catenin and FGF signaling pathways in the interneuromast cells of ak2-deficient embryos. The increased proliferation observed in the interneuromast cells of ak2 null mutants ( Figure 4C) suggest that, in basal conditions, the interneuromast cells seem to be able to initially respond to the Wnt/β-catenin pathway. However, without an external stimulus provided (like AG1478 or 6-BIO treatments which induce a sustained Wnt/β-catenin activation), interneuromast cells fail to generate ectopic intercalary neuromasts. Because it has been shown that the Wnt/β-catenin pathway is upstream of FGF, and that only Fgf signaling is required for subsequent rosette formation, the consistently reduced number and size of rosettes observed in ak2-deficient mutants after AG1478 or 6-BIO treatments suggest a partial impairment of the FGF signaling as well.

ak2 deficiency induces increased cell death in inner ear, primII and PLL neuromasts
Previous studies showed that, in RD patient fibroblasts and zebrafish hematopoietic tissues, ak2 protein can possess an antiapoptotic function (2,8). Notably, for the same samples we also observed a strong signal in the trunk regions between L1 and L2 primary neuromasts, suggesting the presence of apoptotic cells in structures that could represent the primII and the secondary neuromasts ( Figure   6A). To verify this possibility, we repeated the analysis on 4 dpf trunk regions of ak2 mutants in the cldnb:LY-EGFP transgenic background. As shown in Figure 6B

ak2-deficiency triggers oxidative stress response genes
Ak2 deficiency has been linked to an increased level of intracellular ROS and oxidative stress (2,8). In particular, in the hematopoietic tissues of the zebrafish ak2 models of RD we previously described the induction of heme oxygenase 1a (hmox1a) expression (8), which represented a mechanism for cells to defend against potential damage caused by increased oxidative stress(52, 53). The expression of hmox1a and other antioxidant genes is regulated by the binding of the Nrf2 transcription factor to specific enhancer elements known as AREs (or antioxidant response elements)(52, 54).  Figure 7A shows that most of the peroxiredoxin (prdx) genes showed an increased level of expression in the inner ear. Notably, prdx1 expression is also upregulated in several other tissues of the body, including the PLL ( Figure 7A and data not shown). Among the prdx gene family, the only exception was prdx2, which was not expressed in the inner ear and whose expression was reduced in caudal hematopoietic tissue (CHT) and in the primary and secondary neuromasts of the PLL ( Figure 7C). Other examples of genes that were down-regulated in ak2-deficient embryos were the glutathione peroxidase 4 genes (gpx4a and gpx4b); in particular the gpx4b expression appeared strongly reduced in the inner ears ( Figure 7B). The expression of glutathione peroxidase 1a and 1b (gpx1a and gpx1b) and superoxide dismutase 1 and 2 (sod1 and sod2) appeared to be unchanged

Antioxidant treatment partially rescued inner ear hair cell and PLL neuromast defects
Previously, we showed that antioxidant treatment was able to partially rescue the hematopoietic phenotypes observed in the zebrafish models of RD (8). In particular, a treatment with 10-50 µM glutathione reduced the ectopic expression of the oxidative stress marker hmox1a and also restored the expression of c-myb and rag1 (markers of hematopoietic stem cells and lymphoid populations, respectively) in the hematopoietic tissues of ak2-deficient embryos (8). In addition, the treatment of RD patient iPSCs with glutathione (GSH) was able to increase the in vitro differentiation rate of neutrophils (8).
We therefore tested if a similar treatment would be beneficial to the sensorineural defects of a zebrafish RD model. We treated the ak2 hg14 and control embryos with a 2), where we observed a dose dependent response for the rescue that was statistically significant, and the antioxidant treatments were able to rescue the 16%, 25% and 33% of the hair cells, respectively ( Figure 8B and 8C).
Finally, the analysis of the average number of hair cells in the 3 dpf macula of ak2 hg14 mutants and control siblings showed that the macula seems to be less sensitive to the ak2 deficiency with an average of ~20 hair cells (~28 in the controls) which correspond to a 29% of reduction of hair cells, compared to the 48% observed in the cristae of untreated embryos at the same stage or the 69% at 4 dpf. Notably, the GSH treatments were able to almost fully restore the physiological number of hair cells in this specific structure with a 78%, 96% and 94% of rescue. This may indicate that the antioxidant agent might be more effective during the first phases of the ak2 deficiency, suggesting that they might work best when the damage at the target tissue is of a reduced extent.
Unfortunately, because of the anatomical localization of the macula and the consequent difficulties to obtain a good imaging of the structure, we were not able to repeat the analysis at 4 dpf or later, to see if the strong recovery observed at 3 dpf is sustained during time.
We also tested the ability of a GSH treatment to rescue secondary neuromast development. As previously shown, from ~4 dpf secondary neuromasts in ak2 hg14 mutants were atoh1a, atoh1a:dTOM, tnks1bp1:EGFP and pou4f3:GAP-GFP negative ( Figures 3A, 3D, 4C, 5A and 9A). At 5 dpf they usually were also cldnb:LY-EGFP negative and they partially or totally lacked alkaline phosphatase expression ( Taken together, these data indicate that, as previously observed for hematopoietic phenotypes, the administration of an antioxidant treatment (in this specific case GSH) can partially reduce the sensorineural defects observed in an ak2deficient zebrafish model of RD; confirming that these treatments could be potentially helpful in ameliorating a wide range of symptoms in RD patients. 21

Discussion
Reticular dysgenesis is a rare and severe immunodeficiency also characterized by the presence of sensorineural deafness. In the present study, we took advantage of two different zebrafish models of the disease to extend our knowledge of the physiopathology of RD hearing loss, demonstrating that ak2 is expressed in vertebrate sensory organs and that it has a crucial role in zebrafish sensory system development.
The phenotypes observed in our models of ak2 deficiency could explain the mechanism of action resulting in the sensorineural deafness of RD patients. Ak2 deficiency, through increased oxidative stress and cell death, reduces the total number of mature hair cells in the inner ear of mutant animals and it also impairs their regenerative ability.
Moreover, it specifically affects the formation of primary and secondary neuromasts of the posterior lateral line. Finally, we proved that, as previously observed for hematopoietic defects, ameliorating the oxidative stress with an antioxidant treatment partially rescues the sensorineural phenotypes observed in the mutant larvae. However, the lower range of doses required to reduce the oxidative stress and to restore the expression of HSC and lymphoid markers in ak2 null embryos suggest that hematopoietic cells seems to be more sensitive to the oxidative stress than cells from sensory system (and thus easier to rescue with antioxidants alone). This idea seems to be supported by the fact that we never observed sensorineural defects in the hypomorphic ak2 hg16 animals, although they present the same severe hematological phenotypes observed in ak2-deficient embryos (8).
It is interesting to note that the antioxidant treatments presented a different level of efficacy on the two different parts of the zebrafish sensory system. In the inner ear of ak2-deficient mutants the treatment was able to partially rescue the number of mature hair cells (pou4f3-GFP + ) in the cristae and to almost fully restore their correct number in the macula. However, the effect antioxidants had on secondary neuromasts of the PLL was limited to the induction of some of the molecular markers of early neuromasts (atoh1a:dTOM and tnks1bp1:EGFP, cldnb:LY-GFP, and partially alkaline phosphatase) while we never observed the formation of more mature pou4f3:GAP-GFP + hair cells.
Although there are many potential reasons for this observation, one possible explanation could be that it reflects a major difference in the expression profile of hair and supporting cells between primary and secondary neuromasts, with secondary neuromasts relying more heavily on ak2 as opposed to other adenylate kinases encoded in the genome. Alternatively, it could also be due to a distinctive role of ak2 during the different stages of neuromast maturation. In particular, our data seems to suggest that during earlier stages of neuromast development, as observed in secondary neuromasts, ak2 could be involved in maintaining a correct transcriptional profile of the cells possibly via metabolic feedback. Eventually, during the later phases of neuromast development, ak2 role could be more directly linked to the cell survival, as suggested by the increased level of apoptotic cells in inner ear and primary neuromasts. Notably, altered transcriptional profiles have been previously observed in two different in vitro models of RD (10,11). Further investigations will be required to test this hypothesis.
Another possibility is that some pluripotent or multipotent stem cell populations are sensitive to ak2 deficiencies and are incapable of sustained self-renewal, eventually showing stem cell "exhaustion." A variety of mutations have been isolated in RD patients, ranging from missense and nonsense mutations affecting single amino acid positions or the AK2 pre-mRNA splicing, to a range of deletions(1, 2, 4, 5, 28, [59][60][61]. Although the relative impact from the different genomic mutations has not been systematically evaluated, in most of the cases the mutations appear to induce a strong reduction of the AK2 protein level, without significantly affecting the mRNA levels(1, 2, 5, 62). A surprising finding with our RD models was that ak2 hg16 hypomorphic mutants were able to reach adulthood despite the severe hematological defects observed. This hematopoietic phenotype was shared with the ak2 hg14 and ak2 hg15 alleles which were lethal, leading us to believe the mortality in the stronger alleles was not linked to the initial hematopoietic defects (8). WISH analysis showed that ak2 hg16 mutants presented normal levels of ak2 mRNA expression (although protein levels were not determined), while frame-shift mutations in exon 1 in the ak2 hg14 and ak2 hg15 alleles triggered nonsense-mediated decay of ak2 mRNA (8). The lack of defects affecting the sensory systems both during larval and adult phases in ak2 hg16 hypomorphic mutants despite a strong hematopoietic phenotype strongly suggests that, at least in zebrafish, the hematopoietic lineages are significantly more 23 sensitive to ak2 reduction than the sensorineural cells. So far, with only one exception (28), no other major non-hematological defects other than sensorineural hearing loss have been reported in RD patients(5) despite seeing several affected tissues in the zebrafish models of the disease. Identifying all the possible compromised tissues in RD patients will be essential to understanding why bone marrow transplants alone are not sufficient to completely cure the disease.

Zebrafish Lines and maintenance
Zebrafish were maintained and used following protocols approved by the National Human Genome Research Institute Animal Care and Use Committee. All protocols and methods related to animals or animal tissues were approved by animal care and use committee of the National Human Genome Research Institute (G-01-3). Zebrafish handling, breeding, and staging were performed as previously described (63,64). To prevent pigmentation, the embryos used for WISH or confocal analysis were cultured in fish water containing 0.003% 1-phenyl-2-thiouera (PTU, Sigma-Aldrich, cat. # P7629) from 24hpf on. The following strains were used: TAB5 (65)

Whole mount in situ hybridization (WISH)
WISH analysis were performed as previously described (8,69), with minor modifications.
For ak2 antisense probes, the Hybridization Mix was supplemented with 5 % w/v Dextran Sulfate (69). The DIG RNA Labeling kit (SP6/T7; Roche) was used for in vitro transcription using PCR products cloned into the pCR4-TOPO vector (Life Technologies) or directly from PCR fragments (69). All probes were purified using Spin Post-Reaction Clean-Up Columns (Sigma-Aldrich) before use. The following DIGlabeled antisense mRNA probes were generated: eya1, atoh1a, prdx1-6, gpx1a-b, gpx4a-b, sod1-2, gstp1. See Table S1 for a complete list of primers. All embryos used for WISH were fixed overnight in 4% formaldehyde (PFA)/PBS, rinsed with PBS-Tween, dehydrated in 100% methanol, and stored at -20°C until being processed for WISH.
Permeabilization incubation time in 10 µg/ml Proteinase K was optimized to maximize the signal in inner ear or posterior lateral line, with a 1-3 min step depending on the 25 stage (3-5 dpf). Hybridization was performed at 70°C. Stained embryos were stored in 4% PFA until imaging using a ZEISS Axio Zoom V16 stereo microscope.

Histology
After WISH staining, 5-day-old embryos were prepared for embedding in paraffin, transferred to paraffin, oriented, sectioned in ~5 µm transversal sections, and then counterstained using nuclear fast red (Sigma-Aldrich, cat. # N3020). Representative pictures were collected using an upright Zeiss Axio Imager D2 microscope (Carl Zeiss Inc.) with a Plan-Apochromat 20X/0.75 NA or a dry Plan-Neofluar 40X/0.75 NA objective lenses. All images were acquired using a AxioCam HRc full color CCD camera with a 1388 pixel x 1040 pixel imaging field. Zeiss ZEN blue pro 2011 software package was used for collection.

Alkaline phosphatase staining
Alkaline phosphatase staining to visualize the posterior lateral line neuromasts was performed as previously described (34). Briefly, after over-night fixation in 4% PFA, the embryos were washed several times in PBS-Tween and then transferred to WISH staining buffer. The development of the signal was periodically checked under a stereomicroscope and it was stopped by washing with PBS-Tween and finally transferring the embryos in 4% PFA. Stained embryos were stored in 4% PFA until imaging using a ZEISS Axio Zoom V16 stereo microscope.

Inner ear dissection and hair cell bundles staining
Inner ear dissections and phalloidin staining of hair cell bundles was performed as previously described (70). Briefly, after inner ear fixation in 4% PFA the different compartments of the inner ear (saccule, lagena and utricle) were dissected under a stereomicroscope and then stained with Alexa-labeled phalloidin (Alexa Flour 488 Phalloidin, 1:1,000 dilution, Thermo Fisher Scientific) for 30 min at room temperature to visualize hair cell bundles from ak2 hg16/+ and ak2 hg16 fish. Nuclei were stained with DAPI (Thermo Fisher Scientific, cat. #D1306). After the staining, the tissues were washed in PBS-Tween 3 times for 10 min each and then mounted onto a slide with mounting medium. Fluorescent signal was analyzed using a Zeiss LSM 880 confocal system (see below). Quantification was performed selecting representative 2500 µm 2 areas for every structure (6 different areas for utricles, 4 in the case of saccules and lagenas) and the number of phalloidin positive bundles in each area was counted. Then the average number was calculated, and the statistical analysis was performed (see below) (70,71).

Confocal imaging
Confocal images were collected at room temperature using a Zeiss LSM 880 confocal system fitted with an Airyscan module, mounted on an inverted Zeiss Axio Observer Z1 microscope with a Plan-Apochromat 20X/0.75 NA objective lens (Carl Zeiss Inc., Thornwood, NY, USA). Embryos were anesthetized and embedded in 1% low-melting agarose and imaged. All images were acquired in LSM 880 mode using a 32-channel GaAsP-PMT detector. Excitation wavelength of 488nm, 561nm and 405nm lasers were used for green, red and DAPI signals. A range of z-slices at optimal intervals was used depending on the zebrafish orientation to capture all desired structures. Z-sections were collected at defined intervals, and maximum projections were performed on each Zstack. All confocal images were of frame size 512 by 512 or 1024 by 1024 pixels, scan zoom of 0.6 and line averaged 2 times. Zeiss ZEN 2.3 (black) software package was used for collection and post processing of the images (Carl Zeiss).

Hair cell ablation and regeneration assay
Hair cells were ablated at 3, 4, or 5 dpf by treatment with 10 µM copper(II) sulfate

Statistical analysis
The statistical analysis was conducted using two-tailed Student's t-test using Excel (Microsoft) or a one-way ANOVA analysis followed by Tukey's post hoc test (p<0.001) to compare groups of fish treated differently using PRIZM (GraphPad, La Jolla, CA). A difference was considered as significant when the p-value was less than 0.05 for Student's t-test and less than 0.001 for the Tukey's post hoc test. Asterisks refer to significant p-values. Bar graphs showed the mean and the standard errors or the standard error of the mean depending on the test used. All experiments shown were replicated at least two times.                    Time-lapse recording of the migration of primII in a control Tg(-8.0cldnb:LY-EGFP) embryo from ~3.5 to ~4 dpf. Each frame was taken every ~10 min.