A Novel, Ataxic Mouse Model Of Ataxia Telangiectasia Caused By A Clinically Relevant Nonsense Mutation(Part 2)

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3.0 Discussion

By increasing genotoxic stress through the addition of a secondary hit to the DDR pathway, we generated a novel mouse model that displays the most comprehensive set of A-T symptoms of any model to date. This includes severe and progressive ataxia associated with cerebellar atrophy and perturbations of PN properties along with a high incidence of cancer and defects in immune cell development. Together, these comorbidities encompass the three leading causes of premature death in A-T—each contributing to roughly a third of deaths. Of these, the incapacitating effect of ataxia is the most penetrant and is reported by patients and caregivers as having the greatest impact on their quality of life. For this reason, the presence of ataxia and cerebellar atrophy in this new mouse model is of great significance, as it provides for the very first time a resource to not only elucidate the mechanisms of neurological dysfunction, but also a critically needed in vivo model to test severely needed A-T therapeutics, such as the read-through compounds we describe here. We found several similarities between the overall progression of ataxia in the Atm3535; Aptx mice and A-T patients. In clinical A-T, motor deficits are observable by roughly 2 years of age, when parents and doctors detect a lowered ability to transition from toddling to a smooth, reflexively coordinated gate—unfortunately, little is known about motor defects at earlier stages due to the diseases low prevalence and current lack of early diagnostic testing(Rothblum-Oviatt et al. 2016). Patients usually learn to walk without assistance and neurological symptoms tend to remain stable through the first 4 to 5 years of life(Rothblum-Oviatt et al. 2016). We found a similar early progression of motor deficits in Atm735×R3x; Aptx^' mice, detecting early mild motor deficits at P8(righting reflex deficit) followed by a period of relative stability, prior to the onset of progressive and severe ataxia developing after p210 that included changes in gait, startle reflex, tremor, and locomotor activity. Several important questions arise out of these findings, including whether ATM and/or APTX have a neurodevelopmental role in the

cerebellum. Future studies focused on the early phase of the disorder will be critical in understanding if the cerebellum develops normally prior to dysfunction or whether developmental defects are an initial cause. We also found, similar to A-T patients, that the severity of the late-developing ataxia was variable, with some mice ambulating with a clumsy, high-stepping rear gate (Video 3) and others moving almost entirely via contortion of the rear trunk(Fig.1E and Video 4)(Rothblum-Oviatt et al.2016; Levy and Lang 2018; Boder and Sedgwick 1958). Overall, we found that Atm?35×xR35×; Aptx' mice developed a visually profound and measurable progressive loss in motor coordination similar to that observed in A-T patients, which was rescued by expression of at least one copy of the Atm or Aptx get he. The loss of motor coordination in A-T has been attributed to cerebellar degeneration due to its relatively selective neuropathology across the brain and its causal role in several different forms of ataxia(Hoche et al.2012). Consistent with A-T patient neuroimaging studies (Wallis et al.2007; Sahama et al.2015; Sahama et al. 2014; Dineen et al. 2020; Tavani et al. 2003; Quarantelli et al.2013), we find that cerebellar size in Atm735×R3×; Aptx' mice are initially normal, but progressively atrophies concurrently with changes in neurological function. While the loss of cerebellar tissue has been considered a main cause of ataxia in humans, it is unclear from clinical data if ataxia severity is a good predictor of the extent of cerebellar degeneration found postmortem (Aguilar et al. 1968b: Crawford et al, 2006: Dineen et al.2020). In the Atm?35×R35×; Aptx^ mice, we find clear atrophy associated with thinning of the Purkinje neuron dendrite layer that precedes the late, severe behavioral deficits. Our histological observations in the Atm?35xR35×; Aptx'mice suggest that changes in the cerebellar function itself, rather than profound loss of cerebellar cells, are sufficient to cause the ataxic phenotype, consistent with the observation of behavioral defects prior to significant PN loss in several SCAs (Shakkottai et al. 2011:Lorenzetti et al. 2000; Clark et al. 1997; Jayabal et al.2016). The reason why ATM and APTX deficiency is required to generate ataxia in mice when loss of either is sufficient to cause ataxia in humans remains unclear. One possibility is that the rodent brain may more flexibly utilize compensatory pathways or redundant proteins while responding to the 10-20k DNA lesions that impact cells each day(Lindahl and Barnes 2000). Several forms of DNA repair exist to potentially meet this challenge, including base excision repair(BER), nucleotide excision repair (NER),

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as well as homologous and non-homologous end joining (HEJ and NHEJ, respectively), all of which ATM and APTX have been implicated in (Chou et al.2015; Caglayan et al.2017; Wakasugi et al. 2014; Tumbale et al. 2018; Chatterjee and Walker 2017) Alternatively, it may be the case that deficiency in ATM or APTX alone does not adequately impact cell health during the mouse's comparatively short lifespan, and thus eliminating both proteins is necessary to achieve sufficient accumulation of DNA damage to manifest over this time period. This possibility is strengthened by the fact that ATM and APTX have distinct biochemical properties and functional roles in the DNA damage response, and therefore a deficiency in both would be predicted to cause a broader hit to genome stability (i.e., increased genotoxic stress). Our finding that two genome stability pathway proteins are required to induce neurological defects in mice strongly suggests that it is the loss of ATM's Oriole in DNA repair, rather than potential functions in oxidative stress signaling, mitophagy, or mitochondrial function that cause the cerebellar defects (Shiloh 2020). Alternatives, however, cannot be completely ruled out, as APTX, like ATM, has been observed within the mitochondria of brain cells, where it is thought to support the processing of mitochondrial DNA (Meagher and Lightowlers 2014; Sykora et al. 2011). This new mouse model provides a new tool to explore these possibilities and mechanistically define how the loss of ATM and APTX ultimately causes cerebellar dysfunction. The biophysical perturbations observed in PNs recorded from the AtmR35XR35X; Aptx mice are similarly found in several other mouse models of ataxia. This includes changes we observed in PN input resistance, membrane capacitance, and AP threshold and width, which have also been described in mouse models of SCA like 1, 3, and 7 (Stoyas et al.2020; Shakkottai et al.2011; Dell'Orco et al.2015). Moreover, the progressive reduction in PN action potential firing frequency we report positively correlates with the development of ataxia in the AtrnR35XR35X; Aptx'mice is reported in a large number of ataxic mouse models, including SCAs 1, 2, 3,5, 6, and 13 as well as a few episodic forms (see review (Cook, Fields, and Watt 2021)). Given the significant overlap in PN perturbations observed across many different ataxias caused by distinct cellular defects, restoring PN AP firing frequencies has been considered as a broad-based therapeutic approach. However, it remains unclear whether reduced PN firing is a causal factor of ataxia. Moreover, experimental evidence suggests changes in PN activity may in fact be a generalized response to maintain homeostasis during ongoing disease-related impairment of PN physiology (Dell'Orco et al.2015). Thus, continued efforts across all cerebellar ataxias are needed to link the genetic, molecular, and cellular disruptions caused by disease to the specific changes in cerebellar neural signaling that ultimately generate the ataxia. Of significant importance in this effort will be determining whether disease-causing cerebellar defects commonly or differentially cause ataxia through a loss of cerebellar function (e.g., loss of coordinating signals during movement), or from a dominant-negative effect (e.g., disrupting downstream neural circuits with abnormal neural output patterns). Ultimately, while a common therapeutic strategy to address cerebellar ataxias would have the greatest impact, a directed approach that addresses the distinct genetic and molecular causes of cellular dysfunction may ultimately be necessary to successfully develop an efficacious therapeutic.The mechanistic link between a deficiency in DNA stability proteins like ATM and APTX and PN dysfunction is far from clear. Our results suggest the effect of ATM and APTX loss on PNs is intrinsic, as we do not find changes in the presynaptic properties of granule cells or evidence of their cellular loss (no change in GCL thickness). Moreover, while we observed differences in short term plasticity of inferior olivary inputs in ATM- and APTX-deficient PNs and wildtype, these results likely point to a disruption in Ca2* homeostasis potentially via reductions in Inositol 1,4,5-triphosphate receptor 1(ltpr1) expression, similar to those observed in SCAs 1,2, and 3 mouse models as well as ATM-deficient mice (Kim et al.2020; Chen et al.2008; Demirci et al.2009; Shakkottai et al.2011). While this provides a promising avenue for future examination and comparison, it is as of yet unclear, even for the SCAs, whether changes in Ca²* homeostasis， is the causal factor or just another symptom or even compensatory response of diseased or disturbed PNs (Dell'Orco et al.2015). In the immune system, ATM is implicated in the repair of DNA breaks that naturally occur during gene rearrangement of antigen receptor genes in B-and T-cell precursors, a phenomenon critical for antigen receptor(LG and TCR) diversity of these cells.Our finding that T-cell proportions in the blood are significantly reduced is consistent with prior studies in humans and A-T knockout mice(Schubert, Reichenbach, and Zielen 2002; Hathcock et al.2013; Chao, Yang, and Xu 2000; Barlow et al.1996). This reduction of T-cells in the periphery likely correlates with a defect in both cellular and humoral immunity. Importantly, we found that expression of one copy of the ATM gene is enough to restore CD4+ deficits in the blood indicating that therapies able to restore partial ATM expression would have therapeutic efficacy. Although we have not assessed B-cell development in this paper, it is likely that similar conclusions would apply to that process given their mechanistic similarities (Marshall et al. 2018). As expected, the reduction of T-cells in peripheral blood is correlated with defective thymocyte development. In the thymus, we found two main defects. One, induced primarily by APTX deficiency, manifests as a defect in the DN3 to DN4 transition coinciding with early rearrangement of the TCR β locus. The other defect, primarily caused by ATM deficiency, correlates with decreased progression of double-positive CD4*CD8 to single-positive cells, primarily CD4' thymocytes. While the APTX finding was surprising, as its deficiency (AOA 1) is not associated with immune deficits, APTX is known to interact with TCR β gene rearrangement proteins, including XRCC4(Clements et al. 2004). Future studies aimed at defining APTX's role in end-joining mechanisms during TCR gene rearrangement will be

important, and the possibility that alternative end-joining mechanisms, like the use of microhomologies, account for the lack of an immune deficit in its absence needs further investigation (Bogue et al. 1997). The survivability of Atm?35×/R35×; Aptx'mice is considerably longer than prior A-T mouse models. In comparison, the first A-T KO mouse model reported by Barlow et al. died from thymomas usually within 2-4 months after birth (Barlow et al. 1996). The decreased cancer survivability in this and many other knockout A-T mouse models is likely genetic, as the background strain harboring the mutation has been shown to have significant effects on cancer prevalence and survivability, with A/J and C57BL/6

backgrounds having significantly increased survivability over the BALBC and 129S strains(Genik et al. 2014). The fact that our ATM-deficient mice were created on a C57BL/6 background likely underlies; that the Aptx*t mice do not develop ataxia, it is their comparatively long lifespan. Given that the Atm35xR35×; unlikely that the early death in A-T KO mice prevents observation of an ataxic phenotype that would otherwise develop in these mice. However, it is unknown whether the C57BL/6 background confers resilience to developing ataxia, as it does for cancer. Defining the genetic or possibly epigenetic factors that influence the severity of the disease could provide avenues for future therapeutic development. Given the global nature of the ATM and APTX null mutation in our mouse model, we cannot entirely rule out that extra-cerebellar defects may also contribute to the severe ataxic phenotype, and thus future examination outside the cerebellum in the forebrain, brainstem, spinal cord, and even muscle will need to be conducted. Within the cerebellum, while we found some anatomical differences in the PN firing properties within different regions of the cerebellum, we did not detect regional differences in ML width or PN density. However, there are challenges in using regional anatomy as a grouping factor in the cerebellum, as the physical folds of the tissue do not necessarily correlate with the boundaries of functional, molecular expression, or physiological property domains that have been described(Apps and Hawkes 2009; Tsutsumi et al.2015; Gao, van Beugen, and De Zeeuw 2012; Zhou et al.2014). Experiments focused on examining the extent of cerebellar defects within these domains will be important in future studies and compared to the anecdotal reports of anatomical differences in A-T patients(Verhagen et al.2012; De Leon, Grover, and Huff 1976; Amromin, Boder, and Teplitz 1979; Monaco et al. 1988; Terplan and Krauss 1969; Strich 1966; Solitare 1968; Solitare and Lopez 1967; Aguilar et al. 1968a; Paula-Barbosa et al. 1983). While we detect two potential stages in the progression of ataxia in the Atm735wR35x; Aptx mice, the later stage of severe ataxia develops in adulthood in mice, as compared to the childhood-onset in humans. This may limit its use in some neurodevelopmental studies. Also, the interpretation of future experiments must carefully factor in the fact that this new model expresses null mutations in two genome stability genes at the same time, a situation that has not been detected in human patients with either A-T or AOA1. Finally, pinpointing where, when, and how ATM deficiency causes cerebellar pathology and ataxia has been a challenge, as prior ATM-deficient mice generally lack the characteristic features needed to causally link cellular and molecular deficits to the ataxic phenotype. Multiple promising avenues of investigation have been defined, including those focused at the neuronal level, where ATM is implicated in oxidative stress signaling (Chen et al.2003) and synaptic function (Li et al.2009; Vail et al.2016), as well as glial function, where recent evidence suggests glial pathology may be a leading factor in cerebellar pathology(Kaminsky et al. 2016; Campbell et al.2016; Petersen, Rimkus, and Wassarman 2012; Weyemi et al.2015). This novel animal model provides a new tool to test mechanistic hypotheses regarding how ATM deficiency causes cerebellar pathology and ataxia. Additionally, this model may serve most importantly as a critical preclinical tool for testing previously proposed therapeutic candidates (Browne et al.2004; Chen et al.2003) and our own SMRT compounds (Du et al. 2013). The severe limitations of not having a suitable preclinical model for therapeutic testing, especially for a rare disorders like A-T and AOA1, cannot be overstated.

4.0 Materials and Methods

4.1 Ethics Statement

This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All the animals were handled according to approved Institutional Animal Care and Use Committee (IACUC) protocols at The Lundquist Institute (31374-03,31773-02)and UCLA(ARC-2007-082, ARC-2013-068). The protocol was approved by the Committee on the Ethics of Animal Experiments of the Lundquist Institute(Assurance Number: D16-00213). Every effort was made to minimize pain and suffering by providing support when necessary and choosing ethical endpoints.

4.2Mice

All mice were group-housed and kept under a 12-h day/night cycle with food and water available ad libitum. Animals were housed within the general mouse house population, and not in specialized pathogen-free rooms. Older animals were made available wetted food or food gel packs on the ground of the cages as ataxia developed. Atm35 and Atm935×mice were created and provided by Dr. Hicks and colleagues at the University of Manitoba. These mice were created to contain the c.103C>T mutation found in a large population of North African AT patients using recombineering Gateway technology and site-directed mutagenesis. A C>T mutation at this position in the mouse Atm gene creates a TAG G stop codon. The same mutation in the human ATM gene produces a TGA G stop codon. In consideration of the use of these models for therapeutic interventions, we chose to create a mouse model for each of the two PTC codons(Fig.1A). A modified Gateway R3-R4-destination vector was used to pull out the desired region of the mouse Atm gene from a Bacterial Artificial Chromosome (BAC) and subsequently mutated to create either a TAG G stop codon at codon 35(M00001, position 103(C>T))or a TGA G stop codon (M00002, position 103 (CAG>TGA), replicating the human AT PTC). The genomic alleles were then cloned into a modified version of the NorCOMM mammalian targeting vector using a 3-way Gateway Reaction(Bradley et al. 2012). The resulting targeting vectors were electroporated into C2 ES cells(C57BI/6N, derived in A. Nagy lab, Toronto, Canada) and successfully targeted clones were identified by selection with G418 (Gertsenstein et al.2010). Integration of the mutated targeting cassette into the Atm gene locus was confirmed by Southern blot, and by sequencing of PCR products to confirm the presence of the Atm PTC mutation, error-free targeting into the Atm locus, and error-free functional components of the

vector (data not shown). Positive ES clones were used for blastocyst injection to obtain the transgenic lines. The transgenic allele contained a floxed human beta-actin promoter-delta TK1-Neo cassette in the intron upstream of the region containing the mutated exon. This floxed cassette was subsequently excised by crossing with a Cre driver mouse(B6.C-Tg(CMV-are)1Cgn/J) to generate Atm3x+ and AtmM1(103CTMFcc, respectively) mouse lines Atm35×+ (MGI nomenclature: AtmTM1(103CAG>TGA)MFGCc;(Fig.1A). Genotyping of the two Atm lines was performed by using the following primers at Tm 62°℃: Atm gene forward (F) primer:5'-CCTTTGAGGCATAAGTTGCAACTTG-3'; and Atm gene reverse(R)primer: 5'-GTACAGTGTATCAGGTTAGGCATGC-3', creating a wild-type allele product of 151bp or targeted allele product of 241bp (Figs. 1A, 1B).Atmn35~ and Atm035×v were back-crossed with C57Bl/6J mice for 9 generations (99.2% isogenic) prior to cryopreservation and subsequent rederivation using C57BI/6J surrogate mothers. Atm735× and Atm?35xand Atm 35×/Q35× were breeders obtained from F1 sibling Atm35 and Atm835X+ mice. AtmR35×k35×: both were found to be fertile. Aptx knockout (Aptx)mice were created and provided to Dr. Mathews as embryos from Dr. McKinnon (Ahel et al.2006), and subsequently rederived via C57Bl/6J surrogate mothers. Aptx' mice are on a C57Bl/6 and 129 mixed background. Atm35k35; Aptx mice of various wildtype, heterozygous, and homozygous combinations were created from Atm35X; Aptxt breeders were generated by crossing Atm?35×R35 and Aptx^ mice. One cohort of double mutant and corresponding control mice was used in the longitudinal behavioral study for gait analyses and SHERPA testing(Figs.2,3). Multiple additional cohorts of age-matched double mutant and control mice were used for electrophysiological, immunohistological, and Vertical Pole test experiments (Figs.4, 7). Immunological and protein expression experiments were carried out using mice bred from the original AtmR35× and Atm35×rederived mice (Figs.5, 6, and 8). Genotyping was performed from ear tissue samples of P8-11 mice. Real-time PCR methods conducted by Transnetyx Inc. were used to determine each animal's genotype. Animals were made identifiable via toe tattoos given at the same time as ear biopsy. Unique primers for Atm35× and Atm35× were quantified and used to identify wild-type, heterozygous and homozygous mice(listed above). Aptx'and Aptx*" primers were used to assess their genotypes.

4.3 Animal Health

Animals were weighed via a digital scale at P8, 45, 120,210,400. Animal death was recorded as the day found dead, or on the day of euthanization when the animals reached a humane endpoint (animal unable to right itself within the 60s, significant hair matting indicating lack of self-grooming or excessive distress as noted by the veterinary staff). Animal carcasses were immediately frozen upon death, and postmortem necropsies were carried out in batches. The probable cause of death was determined to the best of our ability in collaboration with the staff veterinarian (Dr. Catalina Guerra) by visual inspection of the internal organs. Some mice were cannibalized or accidentally disposed of by vivarium staff and were

therefore labeled as"missing."Mice with no discernable visual cause of death were labeled "indeterminable."Mice that were found with thoracic masses near where the thymus would normally be in young mice were listed as having "thymic cancer."All other identified probable causes of death(e.g., enlarged livers, urinary blockage) were labeled "other."

4.4 Behavior

Before performing any behavioral test, mice were acclimated to the behavioral suite for ~20 minutes. Mice were tested at varying times of the day, in line with their daily cycle. A battery of behavioral tests was performed on naive double mutant mice of the indicated genotypes at various time points depending on the behavior but in the same cohort of mice. The battery of tests included the Catwalk Gait assessment (P45, 120,210, 400) and a subset of the SmithKline-Beecham Harwell Imperial-College and Royal-London-Hospital Phenotype Assessment(SHERPA) tests(P30 and 400). These tests were conducted by the UCLA Behavioral Core. Double mutant and control mice were additionally examined on the Vertical Pole test. All behavioral apparatuses were wiped down with ethanol(70%)between each testing subject.

Gait Analysis

We used a Noldus Catwalk Gait analysis system designed to semi-automatically measure and analyze the gait of mice during normal ambulation. Briefly, the movement of mice across a glass-bottom corridor is video recorded from a central position. Paw prints are highlighted in the video due to light illumination across the glass walking platform. Each mouse step within a video is subsequently detected using Catwalk XT (Noldus) in a semi-automated fashion. A run for each mouse consists of 3 trials of consistent ambulation across the monitored platform. Only consistent trials are accepted, and mice may take up to 10 attempts to complete 3 compliant trials in either direction across the corridor. Compliant trials were defined as those with movement across the platform under 5 s-long and with no more than 60% speed variation. Once placed onto the platform, mice generally ran back and forth without any need for experimenter prompting.

Vertical Pole

Mice are placed at the top of an 80 cm tall bolt with their nose facing down and hind paws as close to the top as possible. Mice are immediately released, and time started immediately upon placement.

Time is stopped when the first forepaw touches the surface below the pole. A mouse's natural predilection is to immediately climb down the pole, and they are given up to 60s to traverse the pole, otherwise, they are helped off the pole. A non-completed trial is automatically given a time of 30s, as 95% of mice that did not descend within 30 s were still on the pole at the 60s mark. SHIRPA Behavioral tests were conducted by the University of California, Los Angeles Behavioral Core at P30

and P400.All parameters are scored to provide a quantitative assessment. which enables the comparison of results both over time and between different laboratories. Each mouse was sequentially tested across all behaviors within ~20-min time span before moving on to the next mouse. The experimenter was blinded to animal genotype. The screen was performed as described previously(Rogers et al. 1997).

Behavioral Observation

The primary screen provides a behavioral observation profile and the assessment of each animal begins by observing undisturbed behavior in a viewing jar(10 cm diameter) for 5 min. In addition to the scored behaviors of body position, spontaneous activity, respiration rate, and tremor, the observer logs any instances of bizarre or stereotyped behavior and convulsions, compulsive licking, self-destructive biting, and retropulsion (walking backward) and indications of spatial disorientation.

Arena Behavior

Thereafter, the mouse is transferred to the arena (30 cm x 50 cm) for testing of transfer arousal and observation of normal behavior. The arena is marked into a grid of 10 x 10 cm² squares to measure locomotor activity within a 30 s-period. While the mouse is active in the arena, measures of startle response, gait, pelvic elevation, and tail elevation are recorded.

Supine Restraint

The animal is restrained in a supine position to record autonomic behaviors. During this assessment, grip strength, body tone, pinna reflex, corneal reflex, toe pinch, wire maneuver, and heart rate Were evaluated.

Balance and Orientation

Finally, several measures of vestibular system function were performed. The righting reflex, contact righting reflex, and negative geotaxis tests were performed. Throughout this procedure vocalization, urination, and general fear, irritability, or aggression were recorded.

Equipment Used

1.Clear Plexiglas arena (approximate internal dimensions 55x33 x18 cm). On the floor of the arena is a Plexiglas sheet marked with 15 squares(11 cm). A rigid horizontal wire (3 mm diameter) is secured across the rear right corner such that the animals cannot touch the sides during the wire maneuver. A grid (40x 20 cm) with 12 mm mesh(approximate) is secured across the width of the box for measuring tail suspension and grip strength behavior. 2. A clear Plexiglas cylinder(15 x ll cm) was used as a viewing jar. 3. One grid floor(40x 20 cm)with 12mm meshes on which viewing jars stand.4. Four cylindrical stainless-steel supports (3 cm length x 2.5 cm diameter) to raise grids off the bench. 5. One square(13 cm)stainless steel plate for the transfer of animals to the arena. 6. Cut lengths of 3/0 Mersilk held in the forceps for corneal and pinna reflex tests 7. A plastic dowel rod was sharpened to a pencil point to test salivation and biting.8. A pair of dissecting equipment forceps, curved with fine points (125 mm forceps, Philip Harris Scientific, Cat. No. D46-174), for the toe pinch. 9.A stopwatch.10. An IHR Click box is used for testing the startle responses. The Click Box generates a brief 20 kHz tone at 90dB SPL when held 30cm above the mouse. Contact Prof. K.P. Steel, MRC Institute of Hearing Research, University Park, Nottingham NG7 2RD. 11.A ruler.12.A 30 cm clear Plexiglas tube with an internaldiameter of 2.5 cm for the contact righting reflex.4.5 Electrophysiology Preparation of acute cerebellar slice cute parasagittal slices of 300 μm thickness were prepared from the cerebellum of experimental and control littermate mice by following published methods (Hansen et al., 2013). In brief, cerebella were quickly removed and immersed in an ice-cold extracellular solution with the composition of (mM): 119 NaCl, 26 NaHCOg,11 glucose,2.5 KCl,2.5 CaCla,1.3 MgCla, and 1 NaHzPOa,pH7.4 when gassed with 5%CO-/95% O2. Cerebella were sectioned parasagittally using a vibratome (Leica VT-100, Leica Biosystems, Nussloch, Germany) and initially incubated at 35°C for～30 min, and then equilibrated and stored at room temperature until use. Extracellular Electrophysiology Extracellular and intracellular recordings were obtained from Purkinje neurons (PNs) in slices constantly perfused with a carbogen-bubbled extracellular solution and maintained at either 37° C (extracellular) or 32°C (intracellular)± 1°C (see above). Cells were visualized with DIC optics and a water-immersion 40× objective (NA 0.75)using a Zeiss Examiner microscope. Glass pipettes of ~3 MQ resistance (Model P-1000, Sutter Instruments, Novato, CA)were filled with extracellular solution and positioned near PN axon hillocks in order to measure action potential-associated capacitive current transients in voltage-clamp mode with the pipette potential held at 0 mV. For whole-cell patch-clamp recordings, pipettes were filled with an intracellular solution(mM): 140 months (CH3KO3S), 10 NaCl, 2 MgCl2，0.2 CaClz，10 HEPES，14 Phosphocreatine （tris salt）， 1 EGTA，4 Mg-ATP，0.4 Na-GTP.100 μM Picrotoxin (Sigma) was added to block inhibitory GABAegeric synaptic inputs. Data were acquired using

a MultiClamp 700B amplifier at 20 or 100 kHz in voltage or current-clamp mode, Digidata 1440 with pClamp10(Molecular Devices, Sunnyvale, CA) and filtered at 2 to 4 kHz. The series resistance was usually between 10 and 15 MQ. Series resistance was compensated at 80% for short-term plasticity experiments only. For extracellular recordings, a total of 20 to 45 PNs were recorded from for each animal across all genotypes, sexes, and age groups. Recordings were distributed across both the medial-lateral and rostrocaudal axis of the cerebellum. Only cells with a "healthy" look(low contrast of cellular borders) and regular, uninterrupted firing rate were examined. During analysis, a few cells were found to have gaps in the firing of greater than 2 seconds, and these cells were eliminated from the analysis, as this type of firing is associated with being "unhealthy."Double mutant tissue did not qualitatively differ in appearance under DIC microscopy prior to recordings, nor was the number of"unhealthy" cells greater than that of other genotypes(7% vs 4 to 11% of all cells across control genotypes at P400). Spatial comparison of neural activity was obtained by recording from serial sections in the flocculus, lateral (2"d or 3'), intermediate (6" or 7th), and medial (11" or 12t)slices. Lower number slices were used in the younger age groups (P45 and 110)to roughly match the relative positioning of recordings across age groups. 0-3 recordings were made from each lobule within each slice dependent on tissue quality and health. Each recording lasted for 1-minute. 3 to 5 mice were used for each age group, and the experimenter was blinded to the genotype, age, and sex.

Intracellular recordings were obtained from PNs in either lobule IIl or VIl of the medial cerebellum(i.e., vermis); no statistical differences in properties were observed between lobules.

Analyses

Spontaneous action potential interstimulus intervals were detected and analyzed using standard and custom routines in ClampFit (Molecular Device), GoPro (Wavemetrics), and Excel (Microsoft). Specifically, action potentials were threshold detected, and spiking statistics(i.e., frequency and interval length)weredeterminedusingadaptedlgorProroutines(TaroTools;https://sites.google.com/site/tarotoolsregister/). The coefficient of variation of the mean inter-spike interval (CV) and the median inter-spike interval(CV2=2 ISIn+1-ISInl/(ISIn+1+ISIn))were calculated in Excel using custom macros. Standard membrane properties were analyzed using lgorPro. RM was determined by averaging 3 voltage trace responses to a-5 mV step pulse from a-80 mV holding potential and measuring the resulting current deflection between 900 and 1000 ms after onset. The membrane time constant was measured by fitting a single exponential to the initial decay phase from 90% to 10% of the peak. CM was calculated by dividing the membrane time constant by the RM.sEPSC events were recorded over a 1-minute epoch and detected and measured using Neuroexpress (v21.1.13). Parallel and climbing fiber axons were stimulated using theta-glass electrodes (W.P.I.) and a TTL-controlled stimulus isolator (ISO-Flex, A.M.P.I.). Evoked EPSC amplitudes and decay time constants (1 exp. for parallel and 2 exp. for climbing fibers) were analyzed using custom routines in lgorPro. Action potentials were examined as part of a set of 1 s current injections between -500 and 2250 pA (250 pA steps) with a holding current adjusted to maintain an ~70 mV potential. Action potential waveforms were measured using custom

routines in lgorPro. Action potential threshold was defined as the first membrane voltage in which the first derivative exceeded 30 mV/ms (Zhu et al.2006).

4.6 Examination of Cerebellar Atrophy

Cerebellar size Immediately after brain removal from the skull, a dorsal, whole-mount image was obtained. Images were then processed using Fiji (NIH). The forebrain and cerebellar sizes were assessed by outlining their 2-dimensional space and then calculating the area. We normalized for possible differences in overall brain size by dividing the results of the cerebellum by forebrain size to produce a relative cerebellum-to-forebrain ratio. Experimenters were blind to the genotype of the animal. Immunohistochemistry At the respective study endpoints (P45, 120,210,400), male and female mice of all genotypes represented in this study was anesthetized with isoflurane and underwent transcardial perfusion with phosphate-buffered saline followed by 4%(w/v) buffered paraformaldehyde (PFA) and then dissected to extract the brain. Images of the whole brain were taken immediately after removing the brain from the skull and the brains were then submerged in 4% PFA for 24 hours, and then cryoprotected in Tris-buffered saline(TBS)with 0.05% azide and 30% sucrose for 72 hours and stored at 4°C until further use. The cerebellum was separated from the forebrain and parasagittal sectioned using a sliding microtome （Microm HM 430， Thermo Scientific） set to section at 40 μm thickness. Cerebellum sections were collected in a series of six and stored in TBS-AF(TBS with 30% sucrose,0.05% sodium azide, and 30% ethylene glycol) at 4°C or -20°C until further use. For immunofluorescent visualization of Purkinje neurons, cerebellum sections of both Atm*; Aptx*t and Atm?35×R35×; Aptx^(n=5 per genotype)were washed for 5 min in TBS three times and then blocked in 15% normal goat serum at room temperature for 30 min followed by free-floating incubation in rabbit or mouse anti-calbindin D-28k (1:1000)for 1 hour at room temperature on an orbital shaker. Sections were then washed for 5 min with TBS three times, followed by free-floating incubation in goat anti-rabbit or mouse Alexa Fluor 488 (1:1000)for 1 h in the dark at room temperature on an orbital shaker. Following secondary antibody incubation, sections were washed for 5 min in TBS three times, then mounted and cover-slipped with Fluoromount-G with DAPI. For some sections, anti-cleaved Caspase-3(1:200) and anti-CD68(1:400) antibodies were additionally probed in parallel with Calbindin using an Alexa Fluor 647 (1:500) secondary antibody. Slides were scanned using Stereo Investigator (MBF Bioscience, ver.2020) on a Zeiss microscope equipped with an ApoTome 2 (Carl Zeiss Microscopy, Axio Imager.M2)using either a 2.5, 10,20,40, or 63x objective, and images captured with a Hamamatsu CMOS camera (Hamamatsu Photonics, ORCA Flash 4.0 LT+).To quantify the number of calbindin-reactive cells in each lobule in the resulting images, we used Stereo Investigator to randomly draw 2 lines between 300 to 500 μm long in each lobule and manually counted the total number of PNs along the length within the 40 um thickness of the tissue slice under 40x magnification.2D density(# of PNs/(linear length*40 um thickness))of the two samples per lobule were then averaged for further comparison between lobules and animals. Calbindin positive PN dendrite widths were measured at a predefined location in lobule VI from each animal in 25 or 40 um thick tissue sections under 20x magnification. Dendritic widths of the primary and secondary branches were measured at the midline between the PN cell bodies and the edge of the molecular layer. Between 7 and 13 dendrites were measured per section, one section per animal. For PN somatic measurements, Stereo Investigator was used to randomly select PNs distributed across the entire medial section under 20x magnification. The average PN width per animal was determined by averaging results across 3 serial sections(16 to 37 PNs per section).PN widths were measured perpendicular to the PN layer or to the exiting dendrite if askew by more than a few degrees. Molecular layer and granule cell layer (visualized with Calbindin and DAPI stains, respectively) widths were assessed in Stereo Investigator by averaging two width measurements at predefined locations for each lobule, roughly halfway along with the long extent of each lobule under 2.5x magnification.CD68 positivity in the cerebellar sections was quantified by measuring the total percent area of CD68+positive staining across the entire medial cerebellar section. 10x stitched images were thresholded to the negative control and quantified using ImageJ, one section per animal. To quantify the percent of Calbindin-positive PNs that were positive for cleaved Caspase-3 we counted PNs across the entire cerebellum using Stereo Investigator. Three, 20x magnification stitched images per animal were examined and the results were averaged. The threshold for Caspase-3 positivity was established from control sections stained with only the secondary antibodies. For non-fluorescent histological analysis, 25-um-thick, free-floating tissue sections onto positively charged slides and air-dried overnight. The tissue was washed in Phosphate-buffered saline(PBS) twice for 5 min, then stained sequentially with 0.1% Hematoxylin in 95% ethanol for~25s and 0.5% Eosin in 95% ethanol for~3s and washed in double-distilled water after each stain. The tissue was subsequently dehydrated for 1 min in 95% ethanol, 100% ethanol, and 100%Xylene washes, then coverslipped with Permount. Slides were imaged using a color camera(Q Imaging, MBF Biosciences) on the same Zeiss microscope and MBF acquisition software. Experimenters were blinded to the mouse genotype in which sections were examined, and the order of examination was interleaved for all histological measurements.

4.7 Flow Cytometry Measurements

Flow cytometry analysis of blood and thymus cells was performed by staining with specific anti-mouse antibodies∶CD4， CD8， CD3， CD44， and CD25. Briefly， whole-blood samples（50 μ） were stained using fluorescent-labeled antibodies, then red blood cells were lysed using BD lysing solution while live white blood cells were stained using a viability stain. Thymi were mechanically dissociated. 1 to 2 million thymus cells were similarly stained using specific antibodies for CD4, CD8, CD44, and CD25. Analysis of immuno-stained white blood cells or thymus samples was performed using FACS ARIA l and data was analyzed using FlowJo software as reported previously (Sanghez et al. 2017).

4.8 Western BIots

Protein extracts (cells/tissues) were homogenized in radioimmunoprecipitation assay (RIPA) lysis buffer (150 mM NaCl,1% Nonidet P-40 [NP-40],0.5% deoxycholate,0.1% SDS,50 mM Tris, pH 8.0) with protease inhibitors(10 ug/ml AEBSF,10 ug/ml leupeptin,5 ug/ml pepstatin,5 ug/ml chymotrypsin,10ug/ml aprotinin). The protein extracts were sonicated and then pelleted by centrifugation at 13,000 rpm for15 min at 4C. BCA protein assay was used to quantify protein concentrations. Samples containing equal amounts of protein 50 to 100 μg per lane were separated using 4 to 12% gradient TGX precast gels BioRad then transferred by TransBlot Semi-Dry BioRad system using Nitrocellulose transfer pack. Transferred blots were stained by Ponceau S stain for equal protein loading then washed and blocked with 5% nonfat dry milk in TBST for 60 min at room temp. Primary antibodies were incubated with shaking overnight at 4°C. Blots were probed for the following antibodies: ATM (D2E2)Rabbit mAb Cell Signaling, at 1:1000 dilution, β-Actin (D6A8)Rabbit mAb Cell Signaling, GAPDH (D16H11)Rabbit mAb Cell Signaling followed by the appropriate horseradish peroxidase-conjugated (HRP)secondary Anti-rabbit, Anti-mouse for 2 hours at room temperature. After multiple washes with TBST, Protein expression was detected by Radiance Plus chemiluminescence substrate using the Azure c400 and the BioRad ChemiDoc imaging systems. Densitometric analysis of the ATM was performed using ImageJ. Experiments were performed with 2 technical and 2-3 biological replicates as indicated.

4.9 Statistical Assessment

The number of animals chosen for each group was based on a priori power analyses using GPower v3.1 based on an α size of 0.5, power of 0.8, and effect sizes estimated from preliminary data or prior studies. We used both parametric (1-and 2-way ANOVA) for normally distributed and non-parametric (Kruskal Wallace) statistical methods for interval data to test for differences between groups followed by pairwise multiple comparisons tests as indicated in the text. Outliers for immune data in Figs. 6 and 7 were excluded via the ROUT method (Q=2%). The specific analyses used for each data set are noted in each figure legend.For all figures: ns not significant,*p≤0.05,** p<0.01,*** p<0.001,**** p<0.0001. Data are reported as mean ± SEM and box and whisker plots indicate the minimum, first quartile, median, third quartile, and maximum data values. All figures and statistical analyses were completed using Excel (Microsoft 360) or Prism v8 and 9 (Graphpad).