A Novel Genomic Strategy Against Hyperuricemia: CRISPR-CAS9-Mediated Restoration of Uricase Function from the Human Pseudogene

Uric Acid: Biochemistry and Evolutionary Context

Uric acid (UA) is the terminal breakdown product of purine metabolism in humans, formed through the sequential oxidation of hypoxanthine to xanthine, and xanthine to uric acid, catalyzed by xanthine oxidoreductase (XOR). In the plasma, uric acid exists predominantly as monosodium urate (MSU) at physiological pH. In most mammals, this relatively insoluble compound is further metabolized to 5-hydroxyisourate—a more soluble intermediate—by the enzyme urate oxidase (uricase, UOX), ultimately yielding allantoin, which is readily excreted in urine. This breakdown process keeps serum uric acid concentrations in non-primate mammals between 1–2 mg/dL, significantly lower than the threshold for crystal formation. A pivotal evolutionary transition occurred approximately 15–20 million years ago during the Miocene epoch, when parallel inactivating mutations rendered the uricase gene (UOX) a pseudogene in the lineage ancestral to modern apes and humans. Two independent nonsense mutations introduced premature stop codons that disrupted the open reading frame, abolishing enzymatic activity. This pseudogenization coincided with a period of climatic cooling, food scarcity, and dietary shifts toward fructose-rich fruits. Several evolutionary hypotheses have been advanced to explain why uricase loss was tolerated or even selected for in ape ancestors: uric acid, as a potent antioxidant, may have compensated for the concurrent evolutionary loss of vitamin C biosynthetic capacity in the Haplorhini clade; elevated UA may have improved cognitive function via enhanced adenosine receptor activity; and elevated UA may have facilitated energy storage during periods of caloric deficit by mimicking insulin resistance and promoting fat deposition from fructose. Regardless of its evolutionary rationale, the loss of uricase has rendered the entire human population constitutively hyperuricemic relative to other mammals. The typical serum uric acid levels for adult men range from 3.4 to 7.0 mg/dL, while for adult women, it ranges from 2.4 to 6.0 mg/dL. These levels approach or exceed the physiological solubility limit of monosodium urate (~6.8 mg/dL at 37°C), creating a chronic state of near-saturated urate in plasma, predisposing individuals to crystal deposition diseases and conferring significant cardiovascular risk.

Epidemiology and Disease Burden of Hyperuricemia

Due to dietary Westernization and high-fructose corn syrup intake, hyperuricemia (serum UA > 6.8 mg/dL by biochemical criteria, or > 7.0 mg/dL clinically in men) has become a global epidemic in recent decades., increased intake of purine-rich foods (red meats, organ meats, shellfish), alcohol (particularly beer), and the rising prevalence of associated metabolic comorbidities including obesity, hypertension, insulin resistance, and chronic kidney disease. According to 2015–2016 National Health and Nutrition Examination Survey (NHANES) data, hyperuricemia affects approximately 20.2% of men and 20.0% of women in the United States. In mainland China, a 2020 meta-analysis reported a prevalence of 13.3% among adults, rising steeply with age and urbanization. Global estimates suggest over 400 million individuals are affected, with incidence continuing to rise worldwide. Clinically, hyperuricemia is the principal etiologic driver of gout, the most common inflammatory arthritis in adults. Gout manifests initially as episodic acute monoarticular arthritis—characteristically affecting the first metatarsophalangeal joint—triggered by MSU crystal deposition in synovial fluid and periarticular tissues, which activates the NLRP3 inflammasome and generates a cascade of interleukin-1β-mediated inflammation. Progression to chronic tophaceous gout involves persistent subcutaneous and periarticular crystal deposits (tophi), destructive arthropathy, and often severe disability. Beyond gout, hyperuricemia is independently associated with a cluster of serious comorbidities: chronic kidney disease (urate nephropathy, nephrolithiasis), cardiovascular disease (hypertension, coronary artery disease, stroke, heart failure), metabolic syndrome, type 2 diabetes, and nonalcoholic fatty liver disease (NAFLD)/nonalcoholic steatohepatitis (NASH). The mechanistic links between uric acid and cardiometabolic disease are multifold. Uric acid activates the NLRP3 inflammasome and stimulates production of pro-inflammatory cytokines. It inhibits endothelial nitric oxide synthase (eNOS), reducing bioavailable nitric oxide and impairing vascular tone regulation—a mechanism directly linking hyperuricemia to hypertension. Uric acid stimulates NADPH oxidase-mediated reactive oxygen species (ROS) production in hepatocytes, adipocytes, and vascular smooth muscle cells, contributing to oxidative stress and endothelial dysfunction. Fatty acid synthase (FAS) and acetyl-CoA carboxylase 1 (ACC1), two lipogenic enzymes that promote triglyceride buildup and steatosis, are upregulated in hepatocytes when intracellular UA triggers the JNK/AP-1 signaling cascade. High uric acid also directly impairs pancreatic beta-cell function through NF-κB-mediated apoptosis and dysregulated insulin secretion, contributing to glucose intolerance. These mechanistic insights, collectively supported by numerous in vitro and in vivo studies—including the Uox-knockout mouse models that spontaneously develop hyperuricemia-associated nephropathy, cardiovascular disease, hepatic steatosis, and metabolic syndrome—strongly support uric acid as a causal driver of cardiometabolic disease, not merely a biomarker.

Current Therapeutic Landscape and Limitations

Contemporary pharmacological management of hyperuricemia rests on two mechanistic pillars: (1) inhibition of xanthine oxidoreductase (XOR) to reduce uric acid synthesis, and (2) augmentation of uric acid excretion or enzymatic degradation. Allopurinol, a purine analog XOR inhibitor discovered by Nobel laureates Gertrude Elion and George Hitchings, remains the first-line therapy globally. Febuxostat is a non-purine inhibitor of xanthine oxidase that provides enhanced selectivity and potency, making it a suitable option for individuals who cannot tolerate allopurinol. Despite widespread use, XOR inhibitors fail to achieve target serum UA levels in approximately 40% of gout patients, particularly those with severe tophaceous disease or renal impairment. Uricosuric agents (probenecid, lesinurad) enhance renal urate excretion but have limited efficacy and are contraindicated in nephrolithiasis. Collectively, these agents address the quantity of uric acid produced but do not resolve the fundamental enzymatic deficiency inherited from primate evolution. Recombinant uricases represent a distinct therapeutic category: by restoring the missing enzymatic step, they can rapidly degrade uric acid to allantoin and dramatically lower serum levels. Rasburicase (Elitek, Fasturtec), a recombinant Aspergillus flavus uricase, is approved only for prevention of tumor lysis syndrome due to extreme immunogenicity with repeated dosing. Pegloticase (Krystexxa), a PEGylated pig-baboon chimeric uricase, was approved in 2010 for refractory chronic gout, but its clinical utility is severely limited: 18% of patients discontinue due to serious adverse events (infusion reactions, cardiovascular events), fewer than 50% achieve sustained serum UA < 6 mg/dL, and up to 40% develop high-titer anti-PEG antibodies that accelerate drug clearance. Ongoing reformulation efforts (modified PEG moieties, co-immunosuppression with methotrexate) and novel delivery systems (enzymosomes, nanozymes, oral uricase) have made incremental improvements but have not fundamentally solved the immunogenicity problem.

This therapeutic gap—the inability to durably, safely, and affordably restore uricase function—motivates the pursuit of genomic engineering approaches. CRISPR-Cas9 technology, by enabling precise, stable genomic integration of a functional uricase gene, offers the prospect of a one-time intervention that permanently restores enzymatic activity in target cells, circumvents recombinant protein immunogenicity, and leverages endogenous cellular machinery for gene expression regulation.

CRISPR-Cas9 Gene Editing: Principles and Therapeutic Applications

The CRISPR-Cas9 system, originally characterized as an adaptive immune mechanism in bacteria and archaea, has been engineered into a powerful and versatile genome editing platform. The system consists of two components: Cas9, an RNA-guided endonuclease containing HNH and RuvC nuclease domains, and a single guide RNA (sgRNA) that directs Cas9 to a complementary genomic target sequence immediately adjacent to a protospacer adjacent motif (PAM, typically NGG for Streptococcus pyogenes Cas9/SpCas9). Upon target binding, Cas9 generates a blunt-ended double-strand break (DSB) at the specified locus. DSBs are subsequently resolved by one of two primary cellular repair mechanisms: error-prone non-homologous end joining (NHEJ), which generates insertions or deletions (indels) disrupting gene function; or homology-directed repair (HDR), which uses a provided donor template to introduce precise sequence modifications, insertions, or transgene knockins.

Precise HDR-mediated knockin, which allows the introduction of functional transgenes, reporter constructs, or correcting sequences at specific genomic sites, is typically chosen for therapeutic purposes over disruptive NHEJ. To enhance HDR efficiency, pharmacological and molecular strategies to suppress NHEJ—including the use of viral proteins such as adenovirus E4orf6/E1B55K that ubiquitinate and degrade DNA ligase IV—have been developed and validated. Safe harbor loci, genomic regions permissive of transgene expression without disrupting endogenous gene function or conferring selective advantage, have been characterized for therapeutic applications. The AAVS1 locus (also known as PPP1R12C intron 1, located on chromosome 19q13), identified as the chromosomal integration site for adeno-associated virus type 2, has emerged as a premier safe harbor for stable transgene expression in human cells, particularly induced pluripotent stem cells (iPSCs) and other therapeutically relevant cell types.

CRISPR-Cas9-based gene editing has demonstrated therapeutic efficacy across numerous genetic diseases. Proof-of-concept studies have corrected mutations causing sickle cell disease, beta-thalassemia, Duchenne muscular dystrophy, hereditary tyrosinemia, and cystic fibrosis. In the oncology space, CRISPR-edited CAR-T cells targeting CD19, CD20, PD-1 knockout T cells, and other immune cell engineering strategies have entered clinical trials. In vivo CRISPR applications have shown efficacy in treating transthyretin amyloidosis, Leber congenital amaurosis, and hereditary angioedema. These clinical advances, combined with continuous improvement in delivery systems (lipid nanoparticles, AAV vectors, ribonucleoprotein complexes) and specificity-enhancing technologies (base editors, prime editors, evolved high-fidelity Cas9 variants), establish CRISPR as a platform technology with broad therapeutic applicability.

CRISPR-Cas9 as a Gene Therapeutic Platform: Current Evidence

The rapid clinical translation of CRISPR-Cas9 gene editing has been remarkable. In 2020, the first in vivo CRISPR therapy—subretinal injection of a Cas9 construct targeting the CEP290 intronic mutation causing Leber congenital amaurosis type 10—was administered (NCT03872479), marking the first direct in-body genome editing in humans. In 2021, Gillmore et al. revealed that in patients with hereditary transthyretin amyloidosis, a single intravenous dosage of lipid nanoparticle-encapsulated Cas9 mRNA and gRNA targeting TTR successfully decreased transthyretin levels by more than 87% with an exceptional safety profile. For hemoglobinopathies, ex vivo CRISPR disruption of the BCL11A erythroid enhancer to reactivate fetal hemoglobin has produced durable, transfusion-independence in patients with sickle cell disease and beta-thalassemia (subsequently culminating in FDA approval of Casgevy/exa-cel in December 2023). These landmark achievements validate CRISPR as a viable therapeutic modality for metabolic and genetic diseases. For hyperuricemia specifically, CRISPR-based approaches have been explored at the preclinical level. The Uox-KO mouse models described above were generated using CRISPR/Cas9, confirming the technology's efficiency for Uox gene disruption. Balico and Gaucher (2021) demonstrated CRISPR-mediated AncUOX knockin into the human AAVS1 locus; this foundational study forms the core focus of the present review and is examined in detail in the sections that follow. These findings collectively support the concept of a CRISPR-based gene therapy for hyperuricemia that leverages the AAVS1 safe harbor for stable, regulated AncUOX expression in human target cells.

Uricase Gene Engineering: Prior Work and Rationale for Genomic Integration

Evolutionary resurrection of the human uricase gene—through progressive mutational reversion of the two inactivating nonsense mutations and restoration of key catalytic residues—has been demonstrated to yield enzymatically active ancestral uricase (AncUOX) proteins with markedly improved stability and activity compared to contemporary fungal or bacterial recombinant uricases. In prior work, Kratzer et al. (2014) demonstrated that ancestral uricase reconstruction via maximum likelihood phylogenetic analysis produced proteins with superior thermostability, pH optimum, and substrate affinity relative to currently approved therapeutics. Furthermore, transient expression of AncUOX in human hepatocytes significantly inhibited fatty acid synthesis, establishing an intriguing link between uricase activity and hepatic lipid metabolism that extends the therapeutic rationale beyond urate-lowering alone. Compared to recombinant protein injection or transitory expression techniques, genomic integration of AncUOX using CRISPR-Cas9 has several advantages: persistent, heritable integration offers sustained enzyme expression; peroxisomal targeting of the AncUOX protein ensures compartmentalized uric acid oxidation with concurrent H₂O₂ neutralization by co-localized catalase; elimination of exogenous recombinant protein administration removes the primary driver of anti-drug antibody development; and the potential for patient-specific cell engineering (autologous transplantation of AncUOX-expressing iPSC-derived hepatocytes or renal tubular cells) offers a personalized therapeutic pathway. In order to assess the data supporting the feasibility of this genome engineering approach, this review looks at the methodical planning, implementation, and characterization of CRISPR-Cas9-mediated AncUOX knockin into the human AAVS1 locus in HEK293 cells as described by Balico and Gaucher (2021).

Molecular Biology of Uricase Pseudogenization and Ancestral Reconstruction

The molecular events underlying uricase gene silencing in the primate lineage have been characterized in detail. Wu et al. (1992) identified two independent nonsense mutations in the human UOX pseudogene: a C→T transition at codon 33 generating a TGA stop codon and a second mutation creating a TAA stop codon at codon 187. These mutations disrupted uricase activity independently in different primate lineages, suggesting convergent evolution toward uricase loss. Additional missense substitutions accumulated over millions of years further diminished residual enzymatic capacity of any UOX gene products. Keebaugh and Thomas (2010) provided a comprehensive evolutionary analysis showing parallel Uox pseudogenization across Haplorhini, elephants, manatees, and certain birds, suggesting that elevated uric acid conferred fitness advantages in specific ecological niches. The ancestral sequence reconstruction approach, pioneered by Kratzer et al. (2014), employed maximum likelihood phylogenetic analysis of UOX sequences from extant mammals to infer the ancestral primate uricase sequence prior to pseudogenization. The reconstructed AncUOX protein, expressed in E. coli, exhibited markedly superior thermostability, catalytic efficiency, and resistance to product inhibition compared to commercial recombinant uricases (rasburicase, pegloticase). AncUOX demonstrated an optimal pH range of 7.4–8.5 and was active at physiologically relevant temperatures and ion concentrations, validating the evolutionary reconstruction approach for generating a superior therapeutic uricase candidate. Significantly, earlier research showed that temporary transfection of human liver cells with constructs expressing AncUOX dramatically decreased the synthesis of fatty acids from fructose, indicating a functional connection between uricase activity and hepatic lipid metabolism that goes beyond urate-lowering.

Uox-Knockout Animal Models and Mechanistic Insights

Murine models of uricase deficiency have provided critical mechanistic insights into the pathophysiology of hyperuricemia. The original Uox-/- mouse generated by Wu et al. (1994) via embryonic stem cell homologous recombination on a hybrid C57BL/6J x 129Sv background exhibited severe urate nephropathy with serum uric acid approximately 12-fold above wild-type controls, but suffered from extremely high neonatal mortality (up to 93%) from renal failure by four weeks of age. A more stable model was generated by Lu et al. (2018) using TALEN-mediated deletion of exon 3, achieving serum UA levels of 420–520 μM with improved but still limited survival. Most recently, Zeng et al. (2024) generated a Uox-/- mouse model via CRISPR/Cas9 deletion of exons 2–4 on a pure C57BL/6J background, producing the most pronounced hyperuricemia (serum UA 1351 μM, 5.5-fold above wild-type) yet reported, but requiring allopurinol rescue to survive beyond 4 weeks. The allopurinol-rescued Uox-/- mice at 8 weeks manifested the full spectrum of hyperuricemia-associated comorbidities: severe urate nephropathy with glomerulosclerosis and tubulointerstitial fibrosis; systolic dysfunction with reduced ejection fraction and stroke volume; aortic endothelial dysfunction with downregulation of ZO-1 and VE-cadherin; hepatic steatosis with oil red O-positive lipid droplets and elevated ALT/AST; and cardiometabolic disturbances including hyperglycemia and hypercholesterolemia. Parallel work using a uricase-deficient rat model (the "Kunming-DY rat," generated by CRISPR/Cas9 deletion of Uox exons 2–4 in Sprague Dawley rats by Yu et al., 2020) demonstrated dramatically better survival (>95% at one year) and moderate hyperuricemia (SUA 48.3 μg/mL in males), providing a more tractable long-term model for studying hyperuricemia-associated pathologies. Both the mouse and rat models confirm the causal relationship between uricase deficiency and the cardiometabolic syndrome, validate therapeutic strategies targeting the uricase pathway, and support the rationale for genomic reintroduction of functional uricase into human cells.

METHODOLOGY

Cell Culture and Maintenance

In the study under review, human embryonic kidney 293 (HEK293) cells were obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA). Cells were maintained in Dulbecco's Modified Eagle Medium (DMEM; Corning, Corning, NY, USA) supplemented with 10% heat-inactivated fetal bovine serum (FBS; Corning) and 1% antibiotic-antimycotic solution (Gibco, Thermo Fisher Scientific, Waltham, MA, USA). All cultures were maintained at 37°C in a humidified incubator with 5% CO₂. Cells were passaged at approximately 80% confluency, approximately three times per week, using 0.25% trypsin-EDTA. All experiments were performed using mycoplasma-tested, low-passage cells (passages 5–15).

Design and Construction of CRISPR Plasmid

According to the original researchers, a parental plasmid (Addgene #64222) encoding mCherry was modified to create the CRISPR expression plasmid (pU6-(BbsI)_CBh-Cas9-T2A-GFP-P2A-Ad4E4orf6). The mCherry coding sequence was removed by restriction digest with FseI and BsrGI (New England Biolabs, Ipswich, MA, USA) and replaced with a PCR-amplified GFP fragment from pX458 (Addgene #48138), enabling visual confirmation of transfection via green fluorescence. The resulting plasmid retains: the human U6 promoter driving sgRNA expression; the CBh promoter driving expression of Cas9-T2A-GFP-P2A-Ad4E4orf6 as a single polyprotein that self-cleaves at T2A and P2A sites; and the adenovirus 4 E4orf6 protein, which forms an E3 ubiquitin ligase complex with E1B55K that ubiquitinates and targets DNA Ligase IV for proteasomal degradation, thereby suppressing NHEJ and enriching HDR-mediated repair outcomes. The authors selected two independent guide RNA sequences targeting the human AAVS1 locus (PPP1R12C intron 1) based on validated published sequences (Chu et al., 2015): gRNA1 (5′-CACCGACCCCACAGTGGGGCCACTA-3′) and gRNA2 (5′-CACCGTCACCAATCCTGTCCCTAG-3′). Complementary oligonucleotides for each gRNA (forward and reverse) were phosphorylated, annealed, and ligated into BbsI-linearized CRISPR plasmid following established protocols. Correct sgRNA insertion was verified by Sanger sequencing using a U6 promoter primer. The gRNA target sites were analyzed for potential off-target cleavage using WTSI Genome Editing), E-CRISP and the CHOPCHOP tool to confirm high on-target specificity with minimal predicted off-target sites in coding regions.

Donor Plasmid Construction

According to the original study, the donor plasmid for AAVS1 knockin was assembled using Gibson Assembly (New England Biolabs) into the backbone of the AAVS1 SA-T2A-puro pA donor (Addgene #22075). from which the puromycin resistance cassette was replaced. The final donor plasmid contained the following elements in order (5′ to 3′): (1) Left homology arm (800 bp), amplified from human HEK293 genomic DNA using primers flanking the upstream AAVS1 sgRNA target site; (2) Splice acceptor sequence from the AAVS1 intron 1/exon 2 junction, enabling the endogenous AAVS1 transcriptional promoter (PPP1R12C) to drive expression of the downstream insert after mRNA splicing; (3) T2A self-cleaving peptide sequence; (4) RFP (red fluorescent protein) coding sequence for FACS-based selection of successful integrants; (5) T2A sequence; (6) AncUOX coding sequence (mammalian codon-optimized, synthesized by GenScript, Piscataway, NJ, USA)—encoding the functionally reconstituted ancestral uricase retaining the conserved C-terminal peroxisomal targeting sequence Ser-Lys-Leu (SKL); (7) Stop codon; (8) SV40 poly(A) signal sequence; (9) Right homology arm (800 bp), amplified from HEK293 genomic DNA flanking the downstream side of the AAVS1 sgRNA target site. The full-length AncUOX coding sequence (codon-optimized for human expression) is 1,017 nucleotides encoding a 338-amino acid protein of approximately 35 kDa. Synthesis and cloning into the donor backbone were performed by GenScript. Bidirectional Sanger sequencing and restriction enzyme analysis were used to confirm each assembly junction. The full nucleotide sequence of AncUOX and translated amino acid sequence are provided (see supplementary materials, Tables S2 and S3 of original publication).

Transfection and Fluorescence-Activated Cell Sorting

In the original study, HEK293 cells were plated at 1 × 10⁶ cells per well in 6-well tissue culture plates 24 hours prior to transfection to achieve 70–80% confluency at time of transfection. Transfections were performed using TransfeX Reagent (ATCC) according to the manufacturer's protocol. For each transfection condition, 2 μg of the gRNA-CRISPR plasmid (either gRNA1 or gRNA2) was combined with 4 μg of the donor plasmid (3:1 molar excess of donor relative to CRISPR vector, by mass approximation) in 250 μL of Opti-MEM reduced serum medium (Invitrogen, Thermo Fisher Scientific) with 12 μL of TransfeX reagent. The DNA-lipid complexes were administered dropwise to cells in full DMEM following a 15-minute incubation period at room temperature to permit lipoplex production. For each guide RNA condition (g1&Donor and g2&Donor), two separate biological replicate transfections were carried out in addition to non-transfected negative controls. GFP fluorescence was first assessed at 72 hours post-transfection by fluorescence microscopy (EVOS FL Auto; Thermo Fisher Scientific) to confirm successful plasmid transfection and Cas9 expression. RFP signal was subsequently monitored, and cells were passaged twice (approximately every 3–4 days) to allow dilution of episomal (non-integrated) donor plasmid before FACS analysis. Bulk cell sorting was performed on a BD FACSAria II flow cytometer (BD Biosciences, San Jose, CA, USA) using a 561-nm laser for RFP excitation. RFP-positive (RFP+) cells were gated using non-transfected controls as negative reference, and RFP+ populations were sorted directly into collection tubes containing DMEM supplemented with 20% FBS and 1% antibiotic-antimycotic to maintain cell viability. Sorted RFP+ cells were then expanded in complete medium until confluent for downstream analyses.

Genomic DNA Extraction and Integration Verification by PCR

The original report states that the DNeasy Blood & Tissue Kit was used to extract genomic DNA from sorted RFP+ cells (both g1&Donor and g2&Donor conditions) and non-sorted wild-type HEK293 controls. QIAGEN, Hilden, Germany) in accordance with the manufacturer's instructions. DNA concentration and purity were assessed by NanoDrop spectrophotometry (A260/A280 ≥ 1.8). Two orthogonal PCR strategies were employed to confirm targeted integration at the AAVS1 locus:

Strategy 1 (Junction PCR, 5′ integration): Primer pair F1 (5′-GGCAGCCTGTGCTGACCCATGCAGTC-3′, annealing ~150 bp upstream of the left homology arm, in the genomic region flanking the 5′ homology arm boundary) and R1 (5′-TAAGATACATTGATGAGTTTGGACAAACCA-3′, annealing in the polyA tract region within the donor insert) was used to amplify a 2,863-bp product spanning from the genomic sequence upstream of the left homology arm through the AncUOX coding region. Amplification of this product is only possible when donor DNA is correctly integrated at the AAVS1 locus, as the forward primer binds genomic sequence absent from the episomal donor plasmid.

Strategy 2 (Internal donor PCR, 3′ integration): Primer pair F2 (5′-AACCCTTACGGCAAGATCACCG-3′, binding within the AncUOX coding sequence) and R2 (5′-CCACAGTTGGAGGAGAATCC-3′, binding in the genomic region downstream of the right homology arm) was used to amplify a 1,201-bp product. This product is specific to correctly integrated donor, as R2 binds genomic sequence not present in the episomal donor plasmid.

All PCR reactions were performed using Q5 Hot Start High-Fidelity Master Mix (New England Biolabs) to minimize amplification errors, with the following conditions: 98°C for 30 seconds (initial denaturation); 35 cycles of 98°C for 10 seconds, 65°C for 30 seconds, 72°C for 2 minutes; 72°C for 2 minutes (final extension). PCR products were analyzed by 1% agarose gel electrophoresis with ethidium bromide staining and GelDoc documentation. Bands of expected size were gel-purified using the QIAquick PCR Purification Kit (QIAGEN) and subjected to bidirectional Sanger sequencing (Eurofins Genomics) to confirm precise integration junctions and absence of indels or point mutations within the AncUOX open reading frame.

Western Blot Analysis

Whole-cell protein lysates from RFP+ (g1&Donor and g2&Donor) cells and non-transfected wild-type HEK293 controls were prepared for Western blot analysis. Cells were washed twice with ice-cold PBS and lysed in denaturing lysis buffer (50 mM Tris-HCl pH 7.5, 1% SDS, 5 mM EDTA, 10 mM β-mercaptoethanol, 1 mM PMSF, 15 U/mL DNase) supplemented with protease and phosphatase inhibitor cocktails (Thermo Scientific). Cell lysates were heated at 95°C for 5 minutes, briefly vortexed, then diluted in NP-40 lysis buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1% NP-40, 5 mM EDTA). Lysates were passed through a 26-gauge needle attached to a 1-mL syringe ten times to shear genomic DNA, incubated on ice for 5 minutes, and clarified by centrifugation at 16,000 × g for 15 minutes at 4°C. The BCA Protein Assay Kit (Pierce, Thermo Scientific) was used to measure the protein concentration. Equal amounts of total protein (40 μg per lane) were resolved by SDS-PAGE on 10% polyacrylamide gels and transferred to PVDF membranes (Bio-Rad, Hercules, CA, USA) at 100 V for 60 minutes. Membranes were blocked in 5% non-fat dry milk in TBST (20 mM Tris-HCl pH 7.6, 150 mM NaCl, 0.1% Tween-20) for 1 hour at room temperature, then incubated overnight at 4°C with primary antibodies: anti-uricase (sc-166070, Santa Cruz Biotechnology, Dallas, TX, USA; 1:500 dilution); anti-catalase (#21260-1-AP, Proteintech; 1:1000); and anti-β-actin (sc-47778, Santa Cruz; 1:1000). Membranes were incubated with HRP-conjugated secondary antibodies for one hour at room temperature following three TBST washes. Chemiluminescent detection was performed using Clarity Western ECL Substrate (Bio-Rad) and imaged with a ChemiDoc Imaging System (Bio-Rad). ImageJ (NIH) has been employed to quantify band intensities.

Immunofluorescence and Confocal Microscopy

To assess AncUOX subcellular localization and confirm peroxisomal targeting, the original investigators performed immunofluorescence colocalization analysis on RFP+ cells (g1&Donor and g2&Donor) and non-transfected control cells. Cells were plated on glass coverslips in 24-well plates at a density of 5 × 10⁴ cells per well and allowed to adhere for 24 hours. Peroxisome labeling was performed using the SelectFX Alexa Fluor 488 Peroxisome Labeling Kit (Invitrogen, Molecular Probes) targeting the peroxisomal membrane protein PMP70, according to the manufacturer's instructions. Briefly, cells were fixed in 4% paraformaldehyde for 15 minutes at room temperature, permeabilized with 0.1% Triton X-100 in PBS for 10 minutes, blocked with 1% bovine serum albumin (BSA) in PBS for 30 minutes, and incubated with anti-PMP70 antibody conjugated to Alexa Fluor 488 (green channel) for 45 minutes at room temperature. Following PMP70 staining, cells were then incubated with anti-uricase antibody (sc-166070, Santa Cruz; 1:200) overnight at 4°C, washed three times in PBS, and incubated with Alexa Fluor 647-conjugated anti-mouse secondary antibody (Thermo Fisher; 1:500) for 1 hour at room temperature. Nuclei were counterstained with DAPI (blue channel; 1 μg/mL in PBS) for 5 minutes. ProLong Diamond Antifade Mountant (Invitrogen) was used to mount coverslips. Confocal images were acquired using a Zeiss LSM 700 laser scanning confocal microscope with a 63× oil-immersion objective (numerical aperture 1.4), using sequential multi-channel acquisition to minimize spectral bleed-through. Using Fiji/ImageJ and the JACoP plugin, images were processed and examined for colocalization in order to get Manders' overlap coefficients and Pearson's correlation coefficient.

Uricase Enzymatic Activity Assay

In the reviewed study, intracellular uricase activity was quantified using a spectrophotometric assay based on the characteristic UV absorption of uric acid at 293 nm. A 1 mM stock solution of uric acid was freshly prepared in 0.1 M sodium phosphate buffer (pH 7.4) by gentle heating and sonication, filtered through a 0.22-μm membrane to remove microcrystals, and confirmed microscopically for absence of substantial crystallization. Working solutions at four concentrations were prepared: 100 μM (physiological), 200 μM (low hyperuricemia, ~1.7 mg/dL), 400 μM (moderate hyperuricemia, ~6.7 mg/dL), and 600 μM (severe hyperuricemia, ~10 mg/dL). RFP+ cells (g1&Donor and g2&Donor) and wild-type control HEK293 cells were plated in 6-well tissue culture plates and grown to 80% confluency. For the assay, cells were washed with PBS and lysed in 1× Assay Buffer from a Uric Acid/Uricase Assay Kit (Cell Biolabs, San Diego, CA, USA) by vigorous trituration and 30-minute incubation on ice with intermittent vortexing. Lysates were clarified by centrifugation at 1,000 × g for 5 minutes at 4°C. The uric acid-containing reaction mixture (400 μL sodium phosphate buffer + 100 μL cell lysate) was incubated at 37°C with gentle agitation, and absorbance at 293 nm was measured using a UV-visible spectrophotometer (NanoDrop or similar) in triplicate at intervals of 0, 1, 5, and 10 minutes. Uricase activity was confirmed by the progressive decrease in A293, reflecting enzymatic oxidation of uric acid substrate. All data were normalized to total protein content as determined by BCA assay. Statistical significance was assessed by two-tailed Student's t-test with p < 0.05 considered significant; p < 0.01 and p < 0.001 were additionally noted.

Statistical Analysis

As reported, all experiments were performed in triplicate (n = 3 biological replicates per condition) unless otherwise stated. Data were presented as mean ± standard error of the mean (SEM). Statistical comparisons between AncUOX-expressing cells (g1&Donor, g2&Donor) and control cells were performed using two-tailed unpaired Student's t-test. For comparisons across multiple time points in the uricase activity assay, one-way ANOVA followed by Tukey's post-hoc test for multiple comparisons was applied. GraphPad Prism 9.0 (GraphPad Software, San Diego, CA, USA) was used for all statistical analyses. Significance thresholds: p < 0.05; p < 0.01; p < 0.001.

RESULT AND DISCUSSION

CRISPR-Cas9 System Design and Genomic Integration Strategy

The overall strategy used by Balico and Gaucher (2021) for CRISPR-Cas9-mediated AncUOX knockin at the AAVS1 locus is illustrated in Figure 1. The human AAVS1 allele at chromosome 19q13 contains a promoter-driven transcript spanning 19 exons. Guide RNA sequences gRNA1 and gRNA2 target sites upstream of exon 2 within the AAVS1 intron 1 region, directing Cas9-mediated DSBs in close proximity to one another. The donor plasmid is flanked by 800-bp homology arms matching genomic sequences immediately surrounding the Cas9 cut sites, enabling high-efficiency HDR-mediated integration. The inserted donor cassette comprises (from 5′ to 3′): a splice acceptor sequence that enables splicing of the endogenous AAVS1 transcript into the donor insert, T2A-RFP (for FACS selection), T2A-AncUOX (the therapeutic payload), and SV40 polyA signal. This architecture leverages the endogenous AAVS1 promoter for transgene expression, ensuring robust and stable transcription without introduction of exogenous promoter sequences that might be subject to silencing. Following successful HDR, the engineered allele contains the complete donor cassette precisely inserted between exon 1 and exon 2 of the AAVS1 locus, as shown in the bottom schematic of Figure 1.

Figure 1. Schematic for CRISPR-Cas9-mediated knockin of AncUOX into the human AAVS1 locus in HEK293 cells. (Top) Wild-type AAVS1 allele with sgRNA1 and sgRNA2 target sites upstream of exon 2. (Center) Donor plasmid architecture with 800-bp homology arms, splice acceptor, T2A-RFP, T2A-AncUOX, and SV40 polyA signal. (Bottom) Engineered allele post-HDR, showing precise integration of the donor cassette between exons 1 and 2 of the AAVS1 locus.

Figure 2. Verification of AncUOX knockin at the human AAVS1 locus. (A) Schematic of engineered allele showing PCR primer positions for junction analysis. (B) Genomic PCR with primers F1/R1 amplifying a 2,863-bp product specific to the 5′ integration junction, present in g1&Donor and g2&Donor cells but absent in controls. (C) PCR with primers F2/R2 amplifying a 1,201-bp product confirming the 3′ integration junction. (D) Western blot of whole-cell lysates showing AncUOX expression (~35 kDa) exclusively in RFP+ cells; catalase and actin serve as loading controls.

Figure 3. Immunofluorescence colocalization analysis demonstrating peroxisomal targeting of AncUOX. Representative confocal images of HEK293 cells stained with DAPI (nuclei, blue), anti-PMP70 (peroxisome marker, green), and anti-AncUOX antibody (red). Merged images show yellow colocalization signal in g1&Donor and g2&Donor cells (rows 2 and 3) but not in non-transfected controls (row 1), confirming functional SKL-mediated import of AncUOX into peroxisomes.

Figure 4. Intracellular uricase activity in AncUOX-expressing HEK293 cells. Time-course measurements of uric acid concentration (μM) in cell lysates of control, g1&Donor, and g2&Donor cells exposed to (A) 100 μM, (B) 200 μM, (C) 400 μM, and (D) 600 μM exogenous uric acid. A decrease in absorbance at 293 nm indicates enzymatic uric acid oxidation. AncUOX-expressing cells demonstrate statistically significant uric acid reduction at all tested concentrations (***p < 0.001; **p < 0.01 vs. control). Data are mean ± SEM from three replicate assays, normalized by total protein concentration.

DISCUSSION:

Implications and Future Perspectives

The findings reviewed here establish, for the first time, that CRISPR-Cas9-mediated stable genomic integration of a reconstructed ancestral uricase gene into the human AAVS1 safe harbor locus confers sustained and enzymatically active uricase function in human kidney cells. These results represent a conceptual milestone: the re-introduction of a metabolic capability that was permanently lost from the primate genome approximately 20 million years ago, accomplished through a single precision editing event that leaves no antibiotic resistance elements, requires no viral integration, and leverages the cell's own PPP1R12C promoter for stable, long-term transgene expression. The therapeutic implications are potentially transformative. Unlike recombinant uricase proteins that are repeatedly administered intravenously and subject to progressive immune neutralization, a genome-integrated AncUOX would be expressed constitutively in the target cell lineage for the cell's lifetime. The peroxisomal targeting of AncUOX addresses the most critical safety concern of exogenous uricase therapy: H₂O₂ toxicity. In the peroxisomal compartment, catalase (present at very high concentrations in most cell types, including hepatocytes and renal tubular cells) rapidly dismutates H₂O₂ to water and oxygen, ensuring that the by-product of uric acid oxidation is safely neutralized. Furthermore, peroxisomal compartmentalization of a foreign protein reduces MHC-I antigen presentation and thus T-cell mediated immune responses compared to cytoplasmic or secreted proteins, potentially enabling sustained AncUOX activity without development of neutralizing antibodies. From the perspective of NAFLD and hepatic lipid metabolism, the findings of Xie et al. (2021) demonstrating that high uric acid induces liver fat accumulation via ROS/JNK/AP-1 signaling, and prior findings that transient AncUOX expression in hepatocytes inhibits fatty acid synthesis from fructose, suggest that AncUOX-expressing liver cells might exhibit reduced lipid accumulation and improved insulin sensitivity in addition to lowered urate levels. Future studies should directly test this hypothesis in AncUOX-expressing hepatocyte cell lines and iPSC-derived hepatocyte organoids exposed to high-fructose or high-purine conditions. The integration of AncUOX into the AAVS1 locus of human iPSCs—which retain pluripotency and can be differentiated into hepatocytes, renal tubular cells, or other urate-metabolizing cell types—represents the most promising pathway toward clinically applicable autologous cell therapy. Important limitations of the reviewed study merit acknowledgment. First, while HEK293 cells are a widely used and tractable model system for proof-of-concept studies, they are transformed embryonic kidney cells rather than primary hepatocytes or renal tubular cells, which are the primary physiological sites of urate metabolism in vivo. The translation of AncUOX knockin to primary hepatocytes, iPSC-derived hepatocytes, or renal tubular epithelial cells remains a critical next step. Second, the study quantified AncUOX activity in cell lysates rather than in intact cells or in vivo, which may not fully recapitulate the peroxisomal compartmentalization dynamics and H₂O₂ management in living cells. Third, off-target editing analysis at the genomic level was not reported in the primary study, and comprehensive next-generation sequencing-based off-target profiling would be essential before clinical translation. Fourth, the relatively modest reduction in uric acid at 600 μM highlights product inhibition as a potential limitation for very severe hyperuricemia, which may require optimization of AncUOX expression levels or co-expression of downstream metabolic enzymes. Fifth, long-term stability of AAVS1 integration and AncUOX expression through multiple cell passages—and potential epigenetic silencing—requires extended characterization.

CONCLUSION

CONCLUSIONS AND FUTURE DIRECTIONS

The evidence reviewed here provides compelling proof-of-concept evidence that CRISPR-Cas9-mediated genomic integration of AncUOX into the human AAVS1 safe harbor locus is technically feasible, results in precisely integrated, sequence-verified constructs, produces enzymatically active, peroxisomally localized uricase protein, and confers measurable uric acid-degrading capacity to human cells across a wide range of clinically relevant uric acid concentrations. These findings represent the first demonstrated genomic re-engineering of the primate uricase pseudogene in human cells. The clinical translational pathway for this technology is clear, albeit requiring substantial further development: (1) AncUOX knockin must be demonstrated in therapeutically relevant cell types—particularly human primary hepatocytes and iPSC-derived hepatocytes, given the liver's central role in urate metabolism and NAFLD pathogenesis; (2) Long-term expression stability and absence of epigenetic silencing at the AAVS1 locus must be confirmed over extended culture periods and through cell passages; (3) Comprehensive off-target editing profiling by unbiased whole-genome sequencing approaches (GUIDE-seq, CIRCLE-seq, or Digenome-seq) is required to establish the safety profile; (4) AncUOX-expressing iPSC-derived hepatocyte organoids should be evaluated in high-urate/high-fructose conditions to test the predicted anti-steatotic effects of restored uricase activity; (5) In vivo efficacy and safety studies in Uox-knockout mouse or rat models should be conducted to assess whether AncUOX cell transplantation or in vivo AAV-CRISPR delivery normalizes serum uric acid, reverses urate nephropathy, and ameliorates cardiovascular and hepatic comorbidities; and (6) Immunological evaluation of AncUOX antigenicity in immune-competent animals is essential to assess whether peroxisomal compartmentalization provides the anticipated immunotolerance advantage. In conclusion, the evidence reviewed here establishes that the evolutionary loss of uricase activity that has burdened the human species with hyperuricemia for 20 million years can be reversed at the genomic level using CRISPR-Cas9 technology. The successful genomic reintegration of AncUOX into human cells, with confirmed protein expression, peroxisomal localization, and enzymatic activity, opens a new frontier in gene therapy for metabolic disease—one that addresses the root evolutionary cause of hyperuricemia rather than merely managing its symptomatic consequences. With continued development, this genomic engineering approach has the potential to provide a durable, safe, and efficacious treatment for millions of individuals worldwide suffering from gout, urate nephropathy, and associated cardiometabolic disorders.

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