Conditional Dystrophin ablation causes profound effects on muscle development, neurobehavior, and extracellular matrix pathways

Conditional Dystrophin ablation in the skeletal muscle and brain causes profound effects on muscle function, neurobehavior, and extracellular matrix pathways | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Conditional Dystrophin ablation in the skeletal muscle and brain causes profound effects on muscle function, neurobehavior, and extracellular matrix pathways View ORCID Profile Muthukumar Karuppasamy , View ORCID Profile Katherine G. English , View ORCID Profile James R. Conner , View ORCID Profile Shelby N. Rorrer , View ORCID Profile Michael A. Lopez , View ORCID Profile David K. Crossman , View ORCID Profile Jodi R. Paul , View ORCID Profile Miguel A. Monreal-Gutierrez , View ORCID Profile Karen L. Gamble , View ORCID Profile Karyn A. Esser , View ORCID Profile Jeffrey J. Widrick , View ORCID Profile Louis M. Kunkel , View ORCID Profile Matthew S. Alexander doi: https://doi.org/10.1101/2025.01.30.635777 Muthukumar Karuppasamy 1 Department of Pediatrics, Division of Neurology at the University of Alabama at Birmingham and Children’s of Alabama , Birmingham, AL 35294 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Muthukumar Karuppasamy Katherine G. English 1 Department of Pediatrics, Division of Neurology at the University of Alabama at Birmingham and Children’s of Alabama , Birmingham, AL 35294 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Katherine G. English James R. Conner 2 Division of Genetics and Genomics, Boston Children’s Hospital , Boston, MA 02115, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for James R. Conner Shelby N. Rorrer 1 Department of Pediatrics, Division of Neurology at the University of Alabama at Birmingham and Children’s of Alabama , Birmingham, AL 35294 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Shelby N. Rorrer Michael A. Lopez 1 Department of Pediatrics, Division of Neurology at the University of Alabama at Birmingham and Children’s of Alabama , Birmingham, AL 35294 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Michael A. Lopez David K. Crossman 3 Department of Genetics at the University of Alabama at Birmingham , Birmingham, AL 35294 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for David K. Crossman Jodi R. Paul 4 Department of Psychiatry and Behavioral Neurobiology, University of Alabama at Birmingham , Birmingham, AL 35294 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jodi R. Paul Miguel A. Monreal-Gutierrez 5 Department of Physiology and Aging, University of Florida , Gainesville, FL 32610, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Miguel A. Monreal-Gutierrez Karen L. Gamble 4 Department of Psychiatry and Behavioral Neurobiology, University of Alabama at Birmingham , Birmingham, AL 35294 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Karen L. Gamble Karyn A. Esser 4 Department of Psychiatry and Behavioral Neurobiology, University of Alabama at Birmingham , Birmingham, AL 35294 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Karyn A. Esser Jeffrey J. Widrick 2 Division of Genetics and Genomics, Boston Children’s Hospital , Boston, MA 02115, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jeffrey J. Widrick Louis M. Kunkel 2 Division of Genetics and Genomics, Boston Children’s Hospital , Boston, MA 02115, USA 6 Department of Pediatrics, Harvard Medical School , Boston, MA 02115, USA 7 The Stem Cell Program, Boston Children’s Hospital , Boston, MA 02115, USA 8 Harvard Stem Cell Institute , Cambridge, MA 02138, USA 9 The Manton Center for Orphan Disease Research at Boston Children’s Hospital , Boston, MA 02115, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Louis M. Kunkel Matthew S. Alexander 1 Department of Pediatrics, Division of Neurology at the University of Alabama at Birmingham and Children’s of Alabama , Birmingham, AL 35294 10 UAB Center for Exercise Medicine at the University of Alabama at Birmingham , Birmingham, AL, 35294 11 UAB Civitan International Research Center (CIRC), at the University of Alabama at Birmingham , Birmingham, AL 35233 12 UAB Center for Neurodegeneration and Experimental Therapeutics (CNET) , Birmingham, AL 35294, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Matthew S. Alexander For correspondence: matthewalexander{at}uabmc.edu Abstract Full Text Info/History Metrics Supplementary material Data/Code Preview PDF Abstract Duchenne muscular dystrophy (DMD) patients suffer from skeletal and cardiopulmonary weakness, and interestingly up to one third are diagnosed on the autism spectrum. Dystrophin is an essential protein for regulating the transmission of intracellular force to the extracellular matrix within the skeletal muscle, but also plays key roles in neurobehavior and cognitive function. The mouse dystrophin gene (also abbreviated Dmd ) is X-linked and has several isoforms with tissue-specific expression, including the large Dp427m muscle transcript found in heart and skeletal muscle, and the Dp427c transcript that encodes the brain-specific dystrophin cerebellar protein. Understanding the functional requirements and pathways that are affected by dystrophin loss will impact dystrophin replacement gene therapy and exon-skipping correction strategies. We generated conditional Dystrophin knockout mice by targeting exon 52 of the mouse Dystrophin ( Dmd flox52 ) locus. We generated dystrophin constitutive and inducible myofiber knockout ( Dmd mKO) mice to evaluate the tissue-specific function of the large skeletal muscle dystrophin isoform. Constitutive embryonic deletion of the Dystrophin gene exclusively in skeletal myofibers resulted in a severe skeletal muscle myopathy, dystrophic histopathology, and functional deficits compared to the mdx mouse. Transcriptomic analysis of skeletal myofibers of the Dmd mKO mice revealed the dysregulation of key extracellular matrix and cytokine signaling pathways. Separately, we generated Purkinje neuron cerebellar dystrophin knockout ( Dmd :Pcp2 KO) mice that displayed neurobehavioral deficits in social approach, social memory, and spatial navigation and working memory. These studies reveal the essential requirement for dystrophin expression in both the skeletal muscle and brain for normal physiological and neurobehavioral function. Significance Statement Duchenne muscular dystrophy is caused by the lack of a functional dystrophin protein in muscle. The large dystrophin (Dp427m) isoform is expressed in skeletal, cardiac, and smooth muscle, but its tissue-specific requirements remain unknown. We generated and characterized a conditional skeletal muscle knockout mouse ( Dmd mKO). Constitutive embryonic genetic ablation of skeletal muscle Dystrophin resulted in muscle histopathologies similar to the mdx mouse, while postnatal muscle Dystrophin ablation resulted in milder pathologies. Ablating cerebellar Dystrophin Dp427c using a Pcp2/L7 -Cre driver resulted in sociobehavioral defects. Transcriptomic analysis of the Dmd mKO mice showed a severe reduction of extracellular matrix and cytokine signaling pathways. Our study reveals an essential role for skeletal muscle dystrophin and identifies essential pathways for modulation using dystrophin-replacement therapies. Introduction In Duchenne muscular dystrophy (DMD) patients there is a disruption of the dystrophin gene (also abbreviated DMD ) that is predominantly expressed in skeletal and cardiac muscle. DMD patients develop progressive muscle weakness in their teenage years that results in severe muscle degeneration and cardiorespiratory complications. The DMD gene is X-linked, thus DMD predominantly affects males who lose the ability to walk by their early teenage years. Mammalian dystrophin protein is an essential protein that is required for normal skeletal muscle function through its key binding to the actin cytoskeleton and dystrophin-associated protein complex (DAPC). When fully sequenced, the human DMD locus was found to be extremely large as it spans 2.2 megabases (Mbs) and has multiple coding regions that are essential for the function of the dystrophin protein( 1 ). The regulation of the dystrophin protein is complex in that the major protein (Dp427) is expressed in multiple tissues and organs including skeletal muscle, heart, lung, intestine, and the brain( 2 ). Further complexity of dystrophin transcriptional regulation exists in that multiple isoforms and DMD promotors exist and the regulation of its expression on a tissue-specific manner has yet to be fully elucidated. The most widely used Duchenne mouse model is the mdx mouse, which harbors an Dmd exon 23 nonsense mutation (C>T) at nucleotide 3185 of the large Dp427m RNA transcript( 3 , 4 ). Several other N-ethyl-N-nitrosourea (ENU) mutagenesis and targeted knockout mouse strains have been generated since the identification of the mdx mouse, that all show varying muscle phenotypes based on the DMD mutation type and Dystrophin exon(s) targeted( 5 ). Additional DMD mouse and other animal models have been generated and assessed for their dystrophin expression, mutation location, muscle pathophysiologies, and their ability to respond to pre-clinical and clinical corrective drugs. Attempts to generate conditional mouse models have had mixed outcomes due the large DMD gene loci and difficulties in targeting the X chromosome using gene editing. One study demonstrated that a dual Dmd exon 2 and 79 floxed mouse when mated with a tamoxifen-inducible cardiac and skeletal muscle MerCreMer transgene resulted in 25-35% dystrophin gene excision( 6 ). A previously homologous recombination (HR)-established knockout mouse targeting dystrophin exon 52 (a known DMD mutation “hot spot” region) resulted in the disruption of the large Dp427, Dp260, and Dp140 dystrophin isoforms( 7 ). This dystrophin exon 52 model ( mdx52 ) displayed severe muscle degeneration pathologies, in addition to cardiac and neurobehavioral defects similar to those observed in the mdx ( Dmd exon 23) mouse( 7 – 10 ). More in-depth comparisons of multiple DMD mouse strains have revealed neurological, cognitive, and behavioral deficits compared to WT controls( 11 , 12 ). Using this knowledge, we generated a conditional-ready Dmd mouse by inserting loxP sites flanking Dystrophin exon 52 ( Dmd flox52 ) that allowed for the ablation of the large Dp427 isoforms upon Cre-mediated excision. Combinatorial mating with skeletal myofiber Cre lines allowed us to define the tissue-specific requirements of the dystrophin protein in skeletal muscle in early and post-natal muscle growth and regeneration. Separate matings with a Purkinje neuronal Cre line also recapitulated several of the learning, cognitive, and social deficits as other DMD mouse lines. Results Generation and initial characterization of skeletal muscle Dystrophin knockout mice We performed CRISPR-targeting of the mouse Dystrophin exon 52 locus and inserted two loxP sequences in the intronic region immediately flanking the exon ( Figure 1A ). Upon Cre-mediated excision of Dmd exon 52, the resulting DNA sequence contains an out-of-frame deletion that impairs the stability of the Dp427, Dp260, and Dp140 transcripts as has been previously shown in the mdx52 mouse( 13 ). We mated male Dmd flox52 /Y mice with the constitutive human skeletal actin ( HSA )-Cre to ablate expression of the large Dp427m protein within the skeletal myofibers ( Figure 1B ). We compared our conditional dystrophin knockout mouse model to the mdx 5cv mouse that has well-characterized dystrophic histological, molecular, and physiological muscle symptoms( 14 ). The resulting Dmd flox52 /Y :HSA-Cre+ ( Dmd :HSA KO) mice have no overt pathologies at birth, but similar to many mdx strains develop a progressive muscle degeneration starting between 4-6 months of age ( Figure 1C ). Analysis of both fast twitch Tibialis anterior (TA) and slow twitch (soleus) muscles from Dmd :HSA KO mice showed increased centralized myonuclei, large areas of fibrosis, and increased collagen production throughout the myofibers ( Figure 1C ). Dmd :HSA KO diaphragm muscle also revealed hallmark muscle degeneration, increased centralized myonuclei, and the presence of collagen deposits compared to Cre-controls ( Figure 1C ). Cross-sectional area (CSA) of Dmd :HSA KO TA myofibers showed an increased frequency of smaller myofibers compared to Cre-controls and similar to those found in our mdx 5cv mice ( Figures 1D and 1E ). Quantitative western blot assessments of Dp427m protein levels in the Dmd :HSA KO TA and soleus muscles showed an average of <5% remaining Dp427m protein in the TA and the soleus muscles ( Figure 1F ). This was further confirmed using immunofluorescent antibody staining in these muscles showing severely reduced levels of the large Dp427m protein ( Supplemental Figure S1 ). Download figure Open in new tab Figure 1. Characterization of a myofiber-specific Dmd conditional knockout mouse. A. Schematic outlining the generation of myofiber-specific Dystrophin conditional knockout mice ( Dmd :HSA KO) by breeding our Dmd flox52 /Y mice to human skeletal actin ( HSA )-Cre transgenic mice. Cre-mediated excision of Dmd exon 52 results in an out-of-frame disruption and instability of the Dp427 RNA transcript. B. Western immunoblotting for the large Dp427m protein in the TA and soleus muscles of WT (open), Dmd flox52 /Y (blue), mdx 5cv (red), and Dmd :HSA KO (green) adult (6-month-old) male mice. C. Densitometry graphs WT, Dmd flox52 /Y , mdx 5cv , and Dmd :HSA KO immunoblots taken from TA and soleus muscle lysates and normalized to Vinculin protein. N = 5 separate mice per cohort. D. Histochemical staining of TA, soleus, and diaphragm muscles isolated from the WT, Dmd flox52 /Y , mdx 5cv , and Dmd :HSA KO male mice. Hematoxylin and eosin (H&E), picrosirius red, and Masson’s trichrome histochemical staining performed. Scale bar = 100 µm. E. Percent fibrosis (%) in the TA muscles of the four mouse cohorts analyzed. F. Percentage (%) centralized myonuclei for the approximately 600 myofibers of the TA muscles of the four mouse cohorts analyzed. G. Cross-sectional area (CSA) displaying myofiber sizes (µm 2 ) shown for each of the four cohorts. Two-way ANOVA with Bonferroni correction performed. P-values of significance are shown: * p < 0.05, ** p < 0.01, *** p < 0.005, and **** p < 0.001. Skeletal myofiber loss of muscle dystrophin greatly impairs muscle physiological function We performed a muscle functional analysis of the Dmd :HSA KO adult mice by performing an assessment of muscle performance following TREAT-NMD standard operating procedures (SOPs) for mdx mice( 15 ). Dmd :HSA KO mice showed reduced rotarod latency to fall, open field test (OFT) total distance travelled, and impaired forelimb grip strength ( Figures 2A - 2C ). All of these locomotor and muscle strength metrics were comparable to those of aged-matched mdx mice. Dystrophin -deficient mice have impaired treadmill running performance due in part to the lack of sarcolemma-associated neuronal NOS (nNOS) binding to Dp427 at the spectrin-like repeats encoded by Dmd exons 45-48( 16 , 17 ). We performed a standardized treadmill downhill running assessment of age-matched male WT, mdx 5cv , Dmd flox52 /Y (Cre negative), and Dmd :HSA KO mice. The Dmd :HSA KO and mdx 5cv mice performed similarly in showing quicker times to exhaustion, reduced overall distance travelled, and reduced velocity compared to internal cohort control mice ( Figures 2D - 2F ). These findings are consistent with the requirement of nNOS-binding to skeletal muscle dystrophin as essential for skeletal muscle exercise and running performance( 18 , 19 ). We assessed physiological force in EDL muscles from our Dmd :HSA KO mice to determine if there were functional deficits in the peak force and resistance to high mechanical strain. Despite their greater mass ( Supplemental Figure S2A ), EDL muscles from the Dmd :HSA KO mice attained similar peak force as EDL’s from the Dmd flox52 /Y Cre-controls ( Supplemental Figure S2B ). This reduction in force per unit physiological cross-sectional area of muscle ( Supplemental Figure S2C ) indicates a reduction in the quality of the Dmd :HSA KO EDL. Eccentric contractions reduced force of muscles from both groups of mice but this response was exacerbated in the Dmd :HSA KO mice ( Supplemental Figures S2D and S2E ). These results suggest that skeletal muscle Dystrophin loss has a moderate reduction in myofiber force output and resistance to mechanical strain in comparison with that of mdx mice. Download figure Open in new tab Figure 2. Adult Dmd myofiber KO mice have a poor running capacity. A. Rotarod analysis shown for the four mouse cohorts: WT, Dmd flox52 /Y , mdx 5cv , and Dmd :HSA KO. B. Grip strength measured for mouse forelimbs shown normalized to grams (g) body weight. C. Open field testing (OFT) of the four cohorts of mice showing distance traveled in centimeters (cm). D. and E. Forced downhill treadmill running of the four cohorts of mice with average velocity and time to exhaustion shown. N = 6 mice/cohort. Two-way ANOVA with Bonferroni correction performed. P-values of significance are shown: * p < 0.05, ** p < 0.01, *** p < 0.005, and **** p < 0.001. Dystrophin myofiber KO mice have impaired skeletal muscle regeneration following injury We performed a cardiotoxin (ctx)-induced injury to the TA muscles of the Dmd :HSA KO and control Dmd flox52 /Y mice to evaluate the regenerative capacity of the skeletal muscle in the absence of the Dp427m protein. The Dmd :HSA KO mice showed a significant amount of fibrosis, collagen, and necrotic regions within the muscle that was comparable to the mdx 5cv strain ( Supplemental Figure S3A ). Immunofluorescent analysis of the myofibers from the Dmd :HSA KO mice showed elevated levels of centralized myonuclei after 21 days post ctx-injury ( Supplemental Figure S3B ). These results demonstrate the absolute requirement for the dystrophin protein in the myofiber and suggest that dystrophin from the muscle satellite cells (MuSCs) cannot compensate for the lack of myofiber dystrophin in the normal regenerative process. Skeletal muscle-specific Dystrophin loss results in cytokine and ECM pathway disruption We performed RNA-sequencing on the Dmd :HSA KO and Cre-control TA muscles to determine what transcript and signaling pathways were disrupted upon the loss of skeletal muscle Dystrophin ( Figure 3A ). Ingenuity Pathway Analysis of RNA transcripts revealed several inflammatory cytokine transcripts that were disrupted as well as extracellular matrix (ECM) including several Collagen isoforms ( Supplemental Figure S4 ). Interestingly, several transcripts known to be disrupted in DMD patients were also disrupted in the Dmd HSA KO mice including an elevation of inducible nitric oxide synthase 2 (iNos ; Nos2 ) ( Figure 3D ). We noted a strong downregulation in key immune-responsive cytokines including Tgfb1 which drives DMD muscle pathologies in Dmd HSA KO mouse muscles and indicates disruption of key cell-cell signaling between the skeletal myofibers and the immune cells residing in the muscle ( Figures 3B - 3D ). These transcriptomic analysis confirms and delineates an essential transcriptomic regulation of key signaling pathways that are known regulators of dystrophin pathophysiology. Download figure Open in new tab Figure 3. Dmd myofiber KO muscles have disrupted cytokine and ECM signaling pathways. A. Volcano plot of the approximately 6,200 dysregulated DEGs in the TA muscles of Dmd :HSA KO mice compared to Dmd flox52 /Y (HSA-Cre-negative) aged-matched littermate controls. B. and C. Heatmaps of significantly dysregulated showing significantly elevated (> 2.0 fold) or downregulated (< 2.0 blue) DEGs with an emphasis on cytokine and ECM pathways. D. RNA-seq DeSeq2 FKPM counts of three candidate highly disrupted transcripts that are validated DMD muscle biomarkers: Nos2 , Col4a4 , and Tgfb1 are shown. Four mice of each cohort ( Dmd flox52 /Y -black; Dmd :HSA KO-red) were analyzed for differentially expressed genes (DEGs). Student’s t-test, two tailed used for comparisons. P-values of significance are shown: * p < 0.05, ** p < 0.01, *** p < 0.005, and **** p < 0.001. Postnatal skeletal myofiber Dystrophin loss impairs muscle maintenance despite dystrophin protein stability We next sought to evaluate the role of skeletal muscle dystrophin in the regulation of adult skeletal muscle maintenance and regeneration. Subsequently, we mated our conditional Dmd flox52 /Y mice to the HSA -MerCreMer tamoxifen-inducible skeletal myofiber Cre line to temporally ablate Dp427m expression in the adult mice ( Figure 4A ). Two months after tamoxifen dosing, we observed 25-30% reduction of total Dp427m protein within the TA skeletal muscles of Dmd :HSA-MerCreMer+ (+ Tam) of Dmd :HSA-MerCreMer+ (-Tam) controls ( Figure 4B ). We observed a similar 20-30% decrease of total Dp427m protein within the gastrocnemius and soleus muscles in the Dmd :HSA-MerCreMer+ (+ Tam) mice ( Figure 4B ). These results are similar to the other reported conditional Dmd mice that showed partial reduction of Dp427mprotein and highlights the stability of the large dystrophin protein after 2 months of protein turnover( 6 ). A histological examination of the Dmd :HSA-MerCreMer+ (+ Tam) mice showed minimal myofiber degradation and small pockets of regenerating myofibers compared to controls ( Figure 4C ). Overall myofiber cross-sectional area (CSA) and minimal Feret diameter analysis revealed smaller clusters of myofibers in the Dmd :HSA-MerCreMer+ (+ Tam) mice compared to no tamoxifen mock controls ( Figures 4D and 4E ). These findings implicate the large amount of stability of the large dystrophin Dp427m protein and implicate protein turnover as an essential part of dystrophin-restoration therapies. Download figure Open in new tab Figure 4. Postnatal loss of muscle dystrophin results in mild skeletal muscle pathologies. A. Schematic for conditional ablation of postnatal dystrophin using a tamoxifen-inducible (± TAM) strategy. Dmd flox52 /Y mice were mated to the HSA -MerCreMer (HSA-MCM) to conditionally ablate dystrophin (+ TAM; red). Dmd :HSA (Cre+) mice given sunflower seed oil (-TAM) were used as controls (black) After a one-week (5x per week) tamoxifen dosing (80 mg/kg body weight) in 8-week-old male mice via intraperitoneal injection, mouse cohorts were evaluated at 0 (pre-dosing), 30, 60, 90, and 180 days (d) post deletion. N = 5 mice per cohort and time period for biopsy. B. Densitometry western immunoblot measurements of the large dystrophin (Dp427m) protein isoform measured in the TA, soleus, and diaphragm muscles. Note the robust persistence of the large dystrophin protein and low protein turnover in the skeletal muscle tissues. C. H&E evaluation of Dmd flox52 /Y :HSA-MCM (Cre+) mice (± TAM) evaluated at 180 days post ablation. Scale bar = 200 µm. D. and E. CSA and minimum Feret diameter measured at 180 days in the TA muscles reveal smaller groupings of myofibers in the md flox52 /Y :HSA-MCM (Cre+) mice (+ TAM) mice compared to (-TAM) controls. Student’s t-test, two tailed used for comparisons. P-values of significance are shown: * p < 0.05, ** p < 0.01, *** p < 0.005, and **** p < 0.001. Loss of cerebellar dystrophin impairs neurobehavioral function and results in ASD-like behaviors Previous studies from our lab and others in various mdx strains demonstrated that the Dp427 cerebellar dystrophin isoform was essential for normal neurobehavior and social development( 12 , 20 – 22 ). We mated the conditional Dmd flox52 /Y mice to L7/Pcp2 -Cre ( Dmd :Pcp2 KO) mice to ablate the Dp427c protein in Purkinje neurons of the cerebellum ( Figure 5A ). Isolated cerebellums from adult Dmd :Pcp2 KO mice showed loss of approximately 65% of cerebellar dystrophin protein compared to Cre- controls ( Figure 5B ; Supplemental Figure S5 ). We performed a series of neurobehavioral assessments in the Dmd :Pcp2 KO mice including social approach and social novelty between novel and familiar objects and mice ( Figure 5C ). Dmd :Pcp2 KO showed impaired social novelty in object recognition compared to Dmd flox52 /Y Cre- control mice ( Figures 5D and 5E ). We performed a similar 3-chamber assessment (novel mouse versus familiar mouse) and the Dmd :Pcp2 KO mice showed no preference for either mouse as compared to controls suggesting that social behavior defects are linked to cerebellar dystrophin protein expression. We performed both T- and Y-maze assessments in the Dmd :Pcp2 KO mice and found no differences in T-maze alternations compared to Cre- control mice ( Figure 5F ). However, in the Y-maze assessment, Dmd :Pcp2 KO mice had reduced alternations without a change in total arm visits which together suggest reduced spatial working memory compared to Cre- controls ( Figure 5G ). Memory tests were also conducted on the Dmd :Pcp2 KO mice that displayed impaired time to find the platform in Morris water maze ( Figure 5H ). These findings support an essential role for cerebellar Purkinje neuronal expression of Dystrophin and demonstrate a functional role for Dp427 in the regulation of neurobehavior in DMD. Download figure Open in new tab Figure 5. Loss of Purkinje neuron brain Dystrophin impairs neurobehaviors in mice. A. Breeding schematic for the Dmd flox52 /Y line to the L7/Pcp2 -Cre (cerebellar Cre driver) transgenic line to generate Dmd:Pcp2 KO mice. B. Densitometry western immunoblot quantification of the large dystrophin (Dp427) protein expression in the cerebellum, cortex, and hypothalamus regions of the brain in the Dmd flox52 /Y (Cre-; open) and Dmd :Pcp2 KO (Cre+; grey) cohorts. N = 5 mice per cohort were analyzed. C. Graphics showing the social approach (novel mouse versus novel object) and social novelty (novel mouse versus familiar mouse) assays performed on the cohorts of mice in a 3-chamber assay system. D. and E. Social approach and social novelty time in chambers (1, 2, and 3) shown between the 3 analyzed mouse cohorts: the Dmd flox52 /Y (open bars), Dmd :Pcp2 KO (grey bars), and mdx 5cv (red bars). Time in chamber and time investigating in minutes (mins) are shown. N = 10 mice per cohort were analyzed with an acclimation period of equivalent amounts. F. and G. T- and Y-maze assessments of the 3 mouse cohorts are shown with spontaneous alternation and the number of total arm visits. H. Morris water maze (MWM) assessments in the 3 cohorts are shown as distance to platform (cm) compared to trial day. Spatial and reverse learning assessments were performed on separate days. N = 10 adult (6-month-old) mice were analyzed for each cohort. Two-way ANOVA with Bonferroni correction performed. P-values of significance are shown: * p < 0.05, ** p < 0.01, *** p < 0.005, and **** p < 0.001. Discussion The generation of novel DMD animal models using CRISPR gene editing and other next-generation genomic DNA editing tools highlights the growing need for comprehensive animal models that accurately reflect disease symptoms and pathophysiologies( 23 ). Our conditional dystrophin knockout model allows for the tissue-specific study of the impact of both dystrophin loss and when combined with dystrophin-replacement strategies, correct DMD strategies. Newer dystrophinopathy mouse models have identified the impact of dystrophin loss on muscle disease symptom progression. The bmx (Becker muscular dystrophy; Dmd exons 45-47 in-frame deletion) mouse has been useful in elucidating the slower muscle disease progression that occurs in patients( 24 ). Our conditional approach allows for the exploration of dystrophin requirements at key developmental and DMD disease-relevant time points. Previous attempts to delineate skeletal muscle dystrophin protein expression using AAV-shRNAi strategies have revealed dystrophin as essential for sarcomeric organization within the muscle myofiber( 25 ). A study of cultured mdx mouse myoblasts revealed a similar downregulation of Collagen and histone transcripts along with impaired fusion and proliferation markers( 26 ). Indeed, there may be a significant amount of epigenetic remodeling that occurs which was shown in embryonic stem (ES) cell cultures of mdx mice and human induced pluripotent stem cells (hiPSCs)( 27 , 28 ). Given the alteration of myogenic determinant transcriptional pathways, consideration of the impact of Dystrophin loss on epigenetic influences on myogenic differentiation is warranted( 29 ). Key autophagy and PTEN/AKT pathways have been shown to be targets for therapeutic amelioration of dystrophic muscle pathologies( 30 – 32 ). Questions remain on the impact and influence of the large dystrophin muscle isoform on muscle satellite cells (MuSCs) and its regulation of cellular polarity( 33 , 34 ). The Dp427 protein is expressed in muscle satellite cells and it is currently unclear if restoration of dystrophin expression in the MuSC population could compensate for myofiber dystrophin loss. Therapies to restore MuSC polarity in DMD MuSCs may be beneficial for restoring the cellular kinetics and polarity defects observed in DMD muscles( 35 , 36 ). It is evident that the expression of dystrophin protein does occur in the brain and both large and smaller dystrophin isoforms likely play independent and critical neuronal functions. Clinical studies have demonstrated that loss of dystrophin is important for normal neurological and neurobehavioral function as up to one-third of all DMD patients are diagnosed on the ASD spectrum with symptoms ranging from mild cognitive impairment to severe developmental delay( 11 , 37 – 39 ). Evaluation of DMD mouse models have shown impaired neurobehavioral responses in memory, learning, and overall electrophysiological neuronal function( 22 , 40 – 42 ). Our Dmd :Pcp2 KO mice show similar defects to those in mdx mice, and highlight the essential role for the large brain dystrophin protein expression. These results argue that newer platforms for exon-skipping drug compounds that can more effectively penetrate the blood-brain-barrier (BBB) may be required for the systemic restoration of dystrophin expression( 43 , 44 ). Our study demonstrates the absolute requirement for dystrophin expression within the skeletal muscle and brain and warrants further evaluation of the tissue-specific pathways impacted by dystrophin loss. Elucidation of these dystrophin-regulated pathways will aide in dystrophin-replacement and correction therapeutic strategies focused on the amelioration of DMD disease pathologies. Materials and Methods Mice Wild type ( C57BL/6J ; strain #000664; Jackson Labs) and mdx 5cv ( C57BL/6J ; strain #002379; Jackson Labs) used in all experiments were maintained in the Alexander lab colony under pathogen-free conditions. Constitutive HSA-Cre ( Acta1 -Cre; strain# 006149) and tamoxifen-inducible HSA-MerCreMer ( HSA -MCM; strain# 031934) were purchased from Jackson Labs (Bar Harbor, ME) and maintained on a congenic C57BL/6J background. The L7/Pcp2 -Cre mice were originally purchased from Jackson Labs and maintained on a congenic C57BL/6J background that had been backcrossed greater than 6 generations (strain #006207; Jackson Labs). All mice had a standard Teklad Global Rodent Diet (Inotiv; West Lafayette, IN) with ad libitum access to food and water. All mouse strains and protocols were approved by the UAB Institutional Animal Care and Use Committee (IACUC) under the protocol 21393. Generation of conditional Dystrophin knockout mice Conditional Dystrophin knockout mice (Dmd flox52 /Y ) were generated commercially (Cyagen; Santa Clara, CA) via CRISPR guide editing of genomic DNA by inserting loxP sites in the intronic regions flanking mouse Dystrophin exon 52 using a 12.2 kb homologous recombination vector. Briefly, loxP sequences 5’-ATAACTTCGTATAGCATACATTATACGAAGTTAT-3’ were inserted in the intronic regions flanking the mouse Dmd exon 52 ( Dystrophin NCBI transcript: NM_007868.6) in C57Bl/6J mouse ES cells (mES). Positively-targeted mouse ES cells were screened for neomycin cassette (later removed via FRT-mediated recombination) and diphtheria toxin subunit A (DTA) selection. After PCR based confirmation of the presence of the loxP sites and removal of the neomyocin cassette, the resultant ES cell line was injected into pseudopregnant females. The subsequent F 0 offspring were genotyped for presence of the loxP sites at mouse Dystrophin exon 52, and then positively-targeted males were mated to WT ( C57Bl/6 ) females to generate F 1 offspring. All positively-targeted F 1 mice were then maintained on the C57Bl/6 strain for subsequent matings. Mice were genotyped for the Dmd flox52 /Y allele using a forward primer “5-TGACTGCATCTGCATACGTG-3” and reverse “5-GAAGCCTGGTAGGCACCAT-3”. Using the Roche Expand Long Template PCR System (MilliporeSigma; Burlington, MA; Cat# 11681842001) the wild type band (2226 bp), targeted loxP conditional allele (2401 bp), and Cre-mediated excised band (992 bp) were used for genotyping following the manufacturer’s recommendations. Later genotyping was performed commercially using Transnetyx probes (Cordova, TN). Isolation of genomic DNA from TA muscles, cerebellum, and tail DNA were performed using Viagen DirectPCR Lysis Reagent (Viagen Biotech; Los Angeles, CA; Cat# 102-T). Histology and Immunofluorescent Imaging Skeletal muscles were slowly cryofrozen as unfixed tissues in Scigen TissuePlus O.C.T. Compound using an isopentane (FisherScientific; Cat# AC397221000) and liquid nitrogen bath as unfixed tissues. Some muscle biopsies were placed in 30% sucrose (MilliporeSigma; Cat# S0389) sink overnight at 4°C prior to the slow cryofreezing to preserve histological and protein integrity. Muscles were sectioned on a Leica CM1950 cryotome at 7-10 µm thickness, and specimens were placed on Fisherbrand Tissue Path Superfrost Plus Gold slides (Fisher Scientific; Cat# FT4981gplus). H&E staining was performed using Hematoxylin (MilliporeSigma; Cat# H9627) and Eosin Y (MilliporeSigma; Cat# 318906) solutions in a previously described protocol( 45 ). Masson’s trichrome staining was performed using a Masson’s Trichrome Stain Kit (Polysciences, Inc; Warrington, PA; Cat# 25088-1) and picrosirius red staining was performed using a Picrosirius Red Stain Kit (Polysciences, Inc; Cat# 24901-500). For immunofluorescent staining, tissue slides were blocked for one hour in a solution of 10% goat serum/1xPBS and incubated for one hour at room temperature using a M.O.M kit (Vector Labs; Cat# BMK-2202; Newark, CA). Tissue slides were set using a VECTORSHIELD Antifade mounting medium with DAPI kit (Vector Labs; Cat# H-1200-10), Immunoblotting Immunoblotting was performed on protein lysates using tissues that were snap frozen in liquid nitrogen and were homogenized using a bead homogenizer and lysing matrix A beads (MP Biomedicals; Cat# 1169100-CF) following the manufacturer’s protocol. Samples were homogenized in T-PER™ Tissue Protein Extraction Reagent (ThermoFisher Scientific; Cat# 78510) supplemented with a protease inhibitor cocktail (ThermoFisher Scientific; Cat# J61852-XF). Protein lysates were quantified using a Pierce BCA Protein Assay Kit (ThermoFisher Scientific; Cat# 23227), and unless otherwise stated, 50 µg of total whole cell lysate was resolved on 4–20% polyacrylamide Mini-PROTEAN ® TGX™ Precast Protein Gels (BioRad). Following an overnight transfer onto PVDF membranes (BioRad; Cat# 1620175), samples were blocked with 5% BSA (RPI Corp.; Cat# A30075) and incubated with primary antibody overnight, washed 3 times, and incubated with secondary antibodies for 1 hour at room temperature. Membranes were washed 4 times for 15 minutes with gentle rocking, before being incubated with SuperSignal™ West Dura Extended Duration Substrate (ThermoFisher Scientific; Cat# 34075). Membranes were imaged on either a ChemiDoc MP Imaging System (BioRad) or a LI-COR Odyssey CLx imager. Antibodies Primary antibodies used in these experiments were validated on dystrophin-deficient tissues or appropriate control tissues for specificity. Primary antisera used for western immunoblotting and immunofluorescent labeling include: dystrophin all isoforms (Abcam; Cat# ab15277), dystrophin Dys1 (Leica; Cat# NCL-DYS1), and vinculin (Cell Signaling Technology; Cat# 7076S). Secondary antibodies used for western immunoblotting included anti-mouse IgG HRP (Cell Signaling Technology; Cat# 7076S) and anti-rabbit IgG HRP (Cell Signaling Technology; Cat# 7074S). Secondary fluorescent anti-mouse and anti-rabbit IgG fluorochrome-conjugated antibodies used include: Alexa Fluor 488 (ThermoFisher Scientific; Cat# A-11001 and A-11008), Alexa Fluor 555 (ThermoFisher Scientific; Cat# A32727 and A32732), and DyLight 633 (ThermoFisher Scientific; Cat# 35563). Fluorescently labeled sections were imaged on a Nikon A1R-HD25 Confocal microscope using NIS Elements software, and later processed using Adobe Photoshop (2025 version). Tamoxifen administration We used a previously-established protocol for the HSA -MerCreMer strain of delivering (80 mg/kg body weight) of (Z)-4-hydroxyl-tamoxifen (4-OHT) (MilliporeSigma; Cat# H7904) mixed with sunflower seed oil (MilliporeSigma; Cat# S5007) via 5 daily intraperitoneal injections while alternating left and right sides to avoid injection site reactions ( 46 , 47 ). Control mice were given equivalent IP injections of sunflower seed oil. Separate mouse cohorts were assessed at 30 day intervals from days 0, 30, 60, 90, and 180 days post injection. Mice were monitored for hepatotoxicity using cytochrome P450 assessments and injection site reactions after administration. Mice locomotor and functional analysis Open field testing and mouse locomotor analysis was measured using a previously established 4 individual chamber sensor system protocol and data was analyzed using Ethovision XT software platform (Noldus; Leesburg, VA)( 48 , 49 ). Mice were analyzed for functional analysis but were first acclimated to the room and open-field chambers one day prior to activity and were given a five minute additional adaptation period prior to activity recording. Mouse locomotor activity was recorded for ten minutes with no external stimulation and the experimenter isolated in a separate room. Physiological force assessments Mice were anesthetized with sodium pentobarbital (80 mg/kg) by intraperitoneal (IP) injection. Isolated EDL muscles were suspended in a phosphate buffer equilibrated with 95% O 2 , 5% CO 2 (35 °C). Muscle contractions were generated using a 150 ms, supramaximal stimulus train (200 µs pulses) with the muscle held at its optimal length (L o ) for tetanic tension. Force was normalized to physiological cross-sectional area assuming a fiber length to muscle length ratio of 0.44 as previously described( 32 ). After evaluation of peak force, muscles were subjected to an active lengthening protocol consisting of the following sequence: one fix-end contraction, 5 lengthening (eccentric) contractions, and two fixed-end contractions. All contractions began with the muscle at L o and were elicited with 150 ms trains at 300 Hz. The lengthening trials consisted of an initial 100 ms fixed-end contraction that allowed the muscle to rise to peak force followed by a constant velocity stretch at four muscle fiber lengths/s to a final length of 120% L o . Force was evaluated at 95 ms of stimulation for both fixed-end and lengthening trials. Social neurobehavior and memory experiments Social approach was determined by placing the test mouse in the middle chamber of a 3-chamber system and determining if the mouse showed preference for a novel mouse or a novel object (round ball). The social approach protocol was adapted from our previous protocol along with other established mouse neurobehavioral assessments in autism spectrum disorder (ASD) mouse models( 20 , 50 ). Mice were evaluated over 10 minute periods after a 10 minute acclimation period in which the novel mouse and novel object were kept in blocked off chambers. Placement of the novel mouse and novel object was switched between the two side chambers between intervals, and the apparatus was cleaned between evaluations. Social novelty was determined by placing the test mouse in the center chamber and evaluating the time spent in chambers containing either an unfamiliar mouse (no previous exposure) or a familiar mouse (aged-matched littermate). The mouse was monitored over a 10 min interval after the 10 minute acclimation period, and its total movement, velocity, time spent in close interaction with familiar or unfamiliar mouse were measured. T-maze and Y-maze (Maze Engineers; Skokie, IL) cohort assessments were performed using previously described with video recordings captured after the mouse was placed in the maze center( 51 , 52 ). Spatial memory and learning evaluation of the mouse cohorts was performed using the Morris Water Maze (Noldus) from an adapted protocol previously used in mdx mice( 53 , 54 ). Experimenter was double-blinded to treatment group and genotype of the mice in each cage and de-blinded occurred after data analysis. Skeletal muscle injury A cardiotoxin-induced injury to the TA was performed by intramuscular injection of 40 µl of 10 µM cardiotoxin (Millipore Sigma; Cat# 217503) given to adult, aged-matched male mice. The contralateral TA muscle was used as a sham control and given an equivalent intramuscular injection of phosphate-buffered saline (1x DPBS; Millipore Sigma; Cat# D8662) given as a control. Mice were evaluated at day 0 (uninjured), 7, 14, and 21 days post cardiotoxin injury for histological changes in muscle architecture and myofiber composition. RNA-sequencing Total RNA was extracted from TA skeletal muscles from aged-matched male mice using TRIzol reagent (ThermoFisher Scientific; Cat# 15596026) for extraction following the manufacture’s protocol. RNA with an RNA Integrity Score (RIN) greater than 7.0 was selected for library amplification. Poly A enrichment using a ribominus kit (ThermoFisher Scientific; Cat# K155002) was performed on the samples. A total RNA library was amplified from approximately 200-300 ng of RNA using a NovaSeq 6000 reagent kit v1.5 (Illumina; San Diego, CA; Cat# 20044417) before being processed on an Illumina NovaSeq 6000 sequencer. Total reads were FASTQ files were uploaded to the UAB High Performance Computer cluster for bioinformatics analysis with the following custom pipeline built in the Snakemake workflow system (v5.2.2)( 55 ). Following count normalization, principal component analysis (PCA) was performed, and genes were defined as differentially expressed genes (DEGs) if they passed a statistical cutoff containing an adjusted p-value =1. Functional annotation enrichment analysis was performed in the NIH Database for Annotation, Visualization and Integrated Discovery (DAVID, v6.8) by separately submitting upregulated and downregulated DEGs. Ingenuity pathway analysis (IPA) was performed on the data as described( 56 ). A p-value <0.05 cutoff was applied to identify gene ontology (GO) terms and the FASTQ experimental files have been uploaded to NCBI’s Gene Expression Omnibus under accession number GSE284723. Quantitative PCR analysis Total RNA was extracted using TRIzol reagent (ThermoFisher Scientific) and the lysing matrix A beads (MP Biomedicals; Cat# 1169100-CF) as described above. Approximately 1 µg of RNA was then reverse transcribed using SuperScript IV First-Strand Synthesis System (ThermoFisher Scientific; Cat# 18091050). Approximately 0.5 µl of sample was then mixed with using TaqMan™ Fast Advanced Master Mix (ThermoFisher; Cat# 4444556) following the manufacturer’s protocol and samples were analyzed on a QuantStudio 5 Real-Time PCR machine (ThermoFisher). The following Taqman real time PCR assays (ThermoFisher Scientific) were used to amplify specific transcripts: Dmd (exon 1, Mm03956078_s1; exons 16-17, Mm00464475_m1; exons 50-51, Mm01216958_m1; exons 64-65, Mm00464520_m1), and Actb1 (Mm02619580_g1). Statistical Analysis For most dual cohort comparisons, either a two-tailed student’s t-test or chi-squared test for normality was performed for all pairwise comparisons. For longitudinal assessments over time or across multiple cohorts, a one-way analysis of variance (ANOVA) with Least Significant Difference (LSD) was performed for all multiple comparisons. GraphPad Prism version 10 software (Graphpad Software; San Diego, CA) was used for all statistical analyses. An a priori hypothesis of *p< 0.05, **p<0.01, ***p<0.001, and ****p< 0.0001 was used for all reported data analyses. All graphs were represented as mean +/- SEM. Acknowledgments and Funding We would like to thank members of the Alexander lab and UAB Center for Exercise Medicine for their critical evaluation of our manuscript. We would also like to thank members of the Kunkel, Gamble, and Esser labs for reviewing our manuscript and assistance with data interpretation. M.S.A. is supported by NIH grants from the Office of the Director under award number (U54OD030167), and an R21 grant from National Institute of Neurological Disorders and Stroke (NINDS) (R21NS140497). Additional funding support was provided by the Hope4Gabe foundation. L.M.K. is supported by NIH National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS) grant R01AR064300, and the Bernard F. and Alva B. Gimbel Foundation. K.L.G. is funded by an NIH National Institute on Drug Abuse (NIDA) grants R01DA046096 and R01NS082413. K.A.E. is funded by NIH NIAMS grants (R01AR066082 and R01AR079220). M.A.L. is supported by an NIH NIAMS grant K08NS120812. Footnotes Competing Interest Statement: L.M.K is a consultant for Pfizer, Dyne Therapeutics, and Myofinity for muscle disease drug therapies. All other authors declare no conflicts of interest. We included Ms. Shelby Rorrer whom was inadvertently omitted. We also edited the wording of the abstract, title, and discussion to reflect the views of the paper more appropriately. https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE284723 References 1. ↵ M. 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