The CRISPR–Cas system serves as an adaptive immune defense mechanism in bacteria and archaea, protecting against invading genetic elements such as bacteriophages and plasmids (28). This system operates through three main stages: adaptation, expression, and interference. During adaptation, foreign DNA fragments, known as protospacers, are incorporated into the CRISPR array as spacers. In the expression stage, the CRISPR array is transcribed into a precursor CRISPR RNA (pre-crRNA), which is then processed into mature crRNAs. Finally, in the interference stage, the mature crRNA directs Cas proteins, which function as endonucleases, to recognize the complementary sequences of the invading genetic element and create DSBs, thereby neutralizing the threat (29, 30).

The CRISPR–Cas system is categorized into two major classes, Class 1 and Class 2, encompassing six types and 33 subtypes. Class 1 is characterized by multiprotein effector complexes and is prevalent in both bacteria and archaea, whereas Class 2, found exclusively in bacteria, relies on a single multifunctional effector protein (31). Each class is further divided into three types: Class 1 includes Types I, III, and IV, whereas Class 2 comprises Types II, V, and VI.

The Class 1 CRISPR–Cas system, particularly Type I, employs multiple Cas proteins that interact with crRNA to form the CRISPR-associated complex for antiviral defense (Cascade). Cascade binds to foreign DNA, allowing crRNA to hybridize with the complementary strand, forming an R-loop, which is then cleaved by Cas3, an enzyme with helicase and nuclease activity (32). The Type I system is the most prevalent among prokaryotes and has further subtypes, such as Type I-A to I-F, each exhibiting distinct Cas protein compositions and mechanisms of interference (33). Similarly, Type III systems, which use Cas10 as the signature protein, exhibit RNA and DNA cleavage capabilities, often functioning through a co-transcriptional targeting mechanism that recognizes RNA and degrades DNA in a coordinated manner (34). Type IV systems, though less characterized, are generally found in plasmids rather than chromosomal CRISPR loci and exhibit a minimal set of Cas proteins, including Cas12a [formerly CRISPR-associated protein 1 (Csf1)]. Their exact function remains under investigation, though they appear to contribute to horizontal gene transfer regulation (35).

In contrast, Class 2 CRISPR–Cas systems rely on single-effector proteins, such as Cas9 in Type II, Cas12 in Type V, and Cas13 in Type VI (36). Cas9, the most well-studied and widely utilized in genome editing, is guided by a single-guide RNA (sgRNA) that directs site-specific DNA cleavage at the protospacer (37). Type II CRISPR systems require a trans-activating CRISPR RNA (tracrRNA) to assist in crRNA maturation and target cleavage (38). Type V systems, which include Cas12 proteins, differ from Cas9 by generating staggered double-strand DNA breaks rather than blunt-ended cuts. Upon specific target DNA recognition, Cas12 undergoes conformational activation that triggers collateral, nonspecific degradation of surrounding single-stranded DNA (ssDNA) molecules. This property forms the mechanistic basis of CRISPR-based diagnostic platforms, where collateral ssDNA cleavage amplifies signal following accurate target detection (39). Type VI systems, featuring Cas13 effectors, uniquely target RNA instead of DNA. After binding to a complementary RNA substrate, Cas13 becomes catalytically activated and induces indiscriminate collateral cleavage of nearby RNA molecules, enabling programmable transcriptome editing as well as highly sensitive RNA diagnostics (40). The diversity of CRISPR–Cas systems highlights their evolutionary innovation and functional specialization in prokaryotic adaptive immunity.

Despite their prevalence in nature and mechanistic diversity, Class 1 CRISPR–Cas systems remain significantly underrepresented in genome-editing applications. Their effector function relies on large, multi-subunit complexes, which introduces major challenges for clinical translation, particularly regarding delivery into mammalian cells, vector packaging constraints, and protein complex assembly fidelity in heterologous systems (32, 41). Additionally, incomplete characterization of many Class 1 subtypes limits precise manipulation and safety assessments, further constraining their therapeutic development compared with Class 2 systems, where single-effector nucleases such as Cas9 and Cas12 are inherently more compatible with current delivery modalities and regulatory expectations (42).

The CRISPR–Cas9 system has revolutionized genome editing due to its high precision and programmability. The breakthrough realization that CRISPR could be repurposed as a genome-editing tool came from the work of Doudna and Charpentier, who demonstrated that the Cas9 protein could be programmed with a synthetic sgRNA to target specific DNA sequences. Their findings laid the foundation for programmable genome modifications, allowing precise genetic alterations in diverse organisms (20, 43). This system consists of two primary components: the Cas9 endonuclease and a guide RNA (gRNA) (44). The Cas9 protein, originally derived from Streptococcus pyogenes (spCas9), is a multidomain endonuclease responsible for generating targeted DSBs in DNA, earning it the designation of a “genetic scissor.” Cas9 comprises two distinct regions: the recognition lobe (REC) and the nuclease lobe (NUC). The NUC region contains the His-Asn-His (HNH) and RuvC nuclease domains, which are responsible for cleaving the complementary and non-complementary DNA strands, respectively, as well as the protospacer adjacent motif (PAM)-interacting domain, which is essential for target DNA recognition (45).

The CRISPR–Cas9 genome editing process follows three key steps: target recognition, DNA cleavage, and repair. The gRNA comprises two essential components: a tracrRNA and a crRNA that guides the Cas9 enzyme to a specific target site within the genome (46). Target recognition occurs when the5′ end of the crRNA hybridizes with a complementary sequence in the genome, while the adjacent PAM sequence facilitates Cas9 binding. Upon successful pairing, Cas9 induces a DSB at the target locus, activating the cellular DNA repair machinery (47). Following DSB formation, cells repair the damage through either non-homologous end joining (NHEJ) or HDR pathways. NHEJ is an error-prone mechanism that directly ligates DNA ends, often introducing insertions or deletions (indels) that can disrupt gene function (48). In contrast, HDR enables precise genome modifications by utilizing a homologous DNA template, facilitating accurate base substitutions or insertions (Figure 1) (49).

Substantial technological refinements have enhanced the specificity and versatility of Cas9. High-fidelity variants such as enhanced specificity Cas9 (eSpCas9) and Streptococcus pyogenes Cas9–High-Fidelity 1 (SpCas9-HF1) have been engineered to minimize off-target cleavage, improving editing accuracy (50). Cas9 nickases (nCas9), which generate single-strand breaks instead of DSBs, further reduce genomic instability while enabling precise editing (51).

Base editors, such as cytosine base editors (CBEs) and adenine base editors (ABEs), enable targeted single-nucleotide modifications, reducing the risk of undesired genomic alterations (52, 53). Base editing is a kind of genome editing that can change a base pair at the target genomic locus directly and irreversibly without the need for donor templates, DSBs, or HDR procedures 41. Two parts make into DNA base editors: a single-stranded DNA modifying enzyme to target nucleotide alteration and a Cas enzyme for DNA binding. There are now two types of DNA base editors available: CBEs and ABEs. Therefore, four transition mutations (cytosine convent into thymine C → T, T → C, guanine into adenine G → A, and A → G) can be installed using base editing (23, 54). Cytosine is changed into uracil by the first base editor (CBE1) deaminating the exocyclic amine. Uracil's detection by cell replication machinery as a thymine, which resulting transition the C–G to T–A (55). Cas proteins are unrelated to gRNA's activity and rely on it for DNA binding. Therefore, by altering their amino acids or changing their structural makeup, one or both of its domains can be rendered inactive with no loss in their ability to bind. By applying this theory, researchers have created modified Cas endonuclease, resulting enable to disrupt the activation of Ruvc and HNH domains to create as nCas or a dCas. All Cas proteins, including dCas and nCas, are able to merge with various other molecules, such as enzymes, without affecting their ability to bind and cut independently (56).

Prime editing, which employs an engineered reverse transcriptase fused to Cas9 nickase, facilitates the precise insertion or deletion of genetic sequences without requiring donor templates, thereby increasing editing efficiency and accuracy (54). The prime editing method was developed by Anzalone in 2019, which decreases the number of inadvertent errors (54, 57). Researchers demonstrated that prime editing effectively installs all 12 potential base-to-base transformations without requiring donor templates of DNA or inducing DSBs in the target sequence, because this method doesn't depend on DSB 46. It consists of a prime editing guide RNA (pegRNA) and Cas9 (H840A) nickase; the pegRNA is longer than gRNA. The engineered reverse transcriptase (RT) results from fusing pegRNA and Cas9 (H840A) nickase. RT serves as the enzyme that uses the pegRNA as a template to synthesize the edited DNA sequence during prime editing. The prime editing has been designed and screened in human cells. There are prime editing systems that process by incorporating Cas9 (H840A) nickase and RT enzyme, and mutations are introduced when transversions occur (58–60). pegRNA includes tracrRNA and spacer, in addition to gRNA, gRNA connected to a specific gene of RNA sequence, which consists of a primer binding site (PBS), the PBS site contains a complementary sequence to the target region of the edited sequence of DNA. Once the target DNA is bound by the CRISPR–Cas9 system, nCas9 cuts the opposing DNA strand to create a single-stranded break (SSB) in the DNA. Using Cas9 fused to RT, the SSB of the DNA sequence links to the PBS site in the sgRNA, acting as a primer to create a new DNA fragment. The targeted DNA will subsequently be replaced by this freshly created DNA (56).

RNA-targeting CRISPR systems (Cas13 and newer RNA effectors) allow for programmable changes to transcripts without changing DNA permanently. This is useful for reversible changes, antiviral strategies, and diagnostics. However, RNA editors have their own problems, such as collateral cleavage in some cases, activity that depends on the sequence, and the need for repeated dosing for therapeutic effects. Newer engineered variants, on the other hand, show better specificity and less collateral activity (61). Table 1 shows some strengths and challenges of contemporary CRISPR editing modalities.

Beyond Cas9, other CRISPR-associated proteins have significantly expanded the potential application. Cas12 (Type V) effectors differ from Cas9 by generating staggered double-strand breaks and exhibiting collateral ssDNA degradation, making them useful for genome editing and diagnostic applications such as DNA endonuclease-targeted CRISPR trans reporter (DETECTR) (62, 63). Cas14/Cas12f and engineered miniaturized variants such as miniaturized CRISPR-associated nuclease (CasMINI) provide compact alternatives suitable for delivery via adeno-associated viruses (AAVs), facilitating in vivo applications where vector size is limiting (64, 65). Cas13 (Type VI) targets RNA rather than DNA, and has been adapted into programmable RNA-editing systems such as RNA editing for programmable A to I replacement (REPAIR) and RNA editing for specific C to U exchange (RESCUE), which catalyze adenosine-to-inosine or cytosine-to-uracil conversions in transcripts, allowing transient, reversible editing without altering the genome (61, 66).

Emerging CRISPR-associated transposase (CAST) systems integrate RNA-guided targeting with transposon machinery, enabling programmable, DSB-free DNA insertions that bypass host repair pathways, thus increasing precision and efficiency for certain genome engineering applications (67).

Finally, CRISPR has been harnessed for gene regulation without altering DNA sequences. CRISPRa employ dCas9 fused to transcriptional activators to upregulate gene expression, while CRISPRi use dCas9-repressor fusions to silence gene transcription, facilitating functional genomics studies and potential therapeutic gene modulation (68, 69). Collectively, these platforms demonstrate the increasing diversification of CRISPR-based technologies, spanning DNA and RNA editing, precision base modifications, delivery system optimization, and transcriptional regulation, thereby expanding their translational potential in next-generation therapeutics.

AD is a progressive ND that impairs cognitive functions, including memory, reasoning, and daily living tasks (70, 71). The disease is a major cause of dementia among the elderly. It is characterized by the presence of amyloid-beta (Aβ) plaques and neurofibrillary tangles in the brain, which impair neurons and contribute to the degeneration of brain tissue (72, 73). The disease initiates with slight memory challenges that progress to severe cognitive and physical impairments, ultimately resulting in complete dependency and premature death (74). The disease disseminates throughout diverse communities at a prominent global health scale due to its elevated prevalence rates. There are 5.5 million individuals in the United States with AD. In 2016, the global number of individuals with dementia reached 43.8 million, reflecting a 116% increase since 1990, according to recent studies (75).

Development of AD largely driven by hereditary elements follows several pathophysiological paths. Many times, mutations in the APP, PSEN1, and PSEN2 genes link fAD to one another (76). The APP gene codes the amyloid precursor protein on chromosome 21, which β-secretase and γ-secretase break to generate Aβ peptides. Applying mutations in APP produces overly high synthesis of Aβ, particularly the Aβ42 variant, which readily accumulates to generate amyloid plaques, a key hallmark of AD (77). Importantly, members of the γ-secretase complex, PSEN1 and PSEN2, respectively encode PSEN1 and PSEN2 proteins (18, 78). Early-onset fAD is caused by mutations in these genes altering γ-secretase activity, hence increasing the Aβ42/Aβ40 ratio and promoting Aβ aggregation (79).

Apart from amyloid pathology, tau protein dysfunction also speeds the development of disease. Although microtubule-associated protein tau (MAPT) gene mutations are rare in AD, tau proteins in affected individuals hyperphosphorylated, therefore losing their ability to stabilize microtubules (80). Along with impairments in neuronal activity, this instability results in paired helical filaments and neurofibrillary tangles (81–83).

Moreover, exacerbating AD development is neuroinflammation. Released by active microglia and astrocytes, reactive oxygen species and pro-inflammatory cytokines damage neurons. Variations in the triggering receptor expressed on myeloid cells 2 (TREM2) gene, which affects microglial activity, have also been associated to increased AD risk, therefore underlining the critical role immune responses play in the evolution of disorders (84–87).

CRISPR–Cas9 represents a formidable gene-editing technique for AD, facilitating the precise modification of pathogenic genes, such as rectifying or eliminating APP, β-site amyloid precursor protein cleaving enzyme 1 (BACE1), and PSEN1/2 mutations to diminish the production and aggregation of harmful Aβ (88); the conversion of Apo-lipoprotein E ε4 allele (APOE4) to the protective APOE3 or APOE2 isoforms, thereby reducing Aβ accumulation, tau pathology, and neuronal susceptibility; the modulation of neuroinflammation through the targeting of CD33, glia maturation factor (GMF), or other immune mediators to enhance microglial clearance of amyloid plaques and inhibit detrimental cytokine signaling (89); and the amplification of neuroprotective pathways by promoting non-amyloidogenic APP processing (α-cleavage) or bolstering microglial functions that maintain neuronal integrity (88). These attributes render CRISPR a versatile method for rectifying genetic disease etiologies, reducing the accumulation of deleterious proteins, mitigating inflammation, and enhancing cerebral resilience. This approach enables researchers to acquire accurate AD models in vivo and in vitro, enhancing the comprehension of disease mechanisms, and aiding in the identification of novel therapy alternatives (Figure 2).

Cell models are extensively utilized in application on many neurodegenerative disorders, including AD, due to the absence of ethical dilemmas, the brevity of the experimental duration, and the cheap associated costs. Over the past few decades, numerous in vitro cellular models of AD have been developed.

Researchers have effectively employed CRISPR–Cas9 to eliminate the APP and BACE1 genes in neural cell types. This resulted in a significant reduction in the synthesis of Aβ, a primary therapeutic target for AD. The previous study demonstrated that the ablation of APP in brain cells via CRISPR–Cas9 diminished both Aβ expression and amyloid pathology. This indicates its potential as a gene therapy alternative for individuals with APP mutations (89, 90). Sun et al. (91) employed CRISPR–Cas9 to modify the C-terminus of endogenous APP, markedly diminishing β-secretase cleavage and redirecting processing toward non-amyloidogenic α-secretase, thus decreasing Aβ generation.

Previous studies were conducted to reduce the Aβ utilizing CRISPR technique the knockout of PSEN1 and PSEN2 genes (92–95). In addition, researchers effectively introduced this mutation into brain cells utilizing Cas9 nickase–deaminase, significantly reducing Aβ generation (96). Simultaneously, Song et al. (97) discovered that the suppression of KIBRA (Kidney and Brain expressed protein) in HT22 cells inhibited proliferation and induced death, whereas treatment with Aβ1–42 oligomers was administered (96, 98). Prime-Base editing was used to treat AD via utilizing CRISPR to correct the gene defects, and the results shows reduction in Aβ peptide accumulation. Several studies have been performed in vitro utilizing CRISPR–Cas9 technology to modify and editing the APOE gene, as this method has the potential to convert high-risk APOE4 alleles into protective alleles such as APOE3 or APOE2 to reduce the AD risk (99).

In addition to constructing AD cell models, CRISPR–Cas9 technology can also be utilized to develop AD animal models to improve and correct the cognitive function in AD. Serneels, T'Syen (100) Developed a novel model utilizing the CRISPR–Cas9 technique to construct a humanized A sequence in the APP gene of both species (100). Another study utilized animal models in vivo to improve cognitive function and correct APP and BACE1 genes in AD. It demonstrated that the CRISPR system can suppress Aβ, having a significant effect on both pathology and cognitive function (101–103).

Furthermore, CRISPR-based gene editing techniques, particularly base editing and prime editing, exhibit the capacity to rectify harmful AD mutations without inducing double-strand DNA breaks. The genetic modification executed by base editors led to significant enhancements in tau pathology and cognitive recovery in the examined mice (58, 104, 105). APOE possesses various variations, including APOE2, APOE3, APOE3r, and APOE4. Komor et al. (59), the transfer of APOE3 to APOE4 in mouse astrocytes was achieved by changing Cincodon158 to T using the CRISPR–Cas9 system, indicating that point mutations were corrected by the CRISPR–Cas9 approach (59).

Prime-Base editing serves as a genome-editing technique that offers extensive genetic modification possibilities via mutation repair without inducing DSBs to address AD. In vivo research focused on AD-related applications needs greater exploration (104–106), such as a study by Park et al. (103) via utilizing Cas9 nanocomplexes with sgRNAs targeting the tyrosine hydroxylase and BACE1 genes were administered into mouse primary neural cells to evaluate the efficacy of Cas9 nanocomplexes. Furthermore, Takalo et al. (107) utilizing the CRISPR–Cas9 gene editing technique and evaluating the preventive effects of the Plc2-P522R variant.

Patient-derived induced pluripotent stem cells (iPSCs) have garnered growing interest due to their distinctive physiological characteristics. AD mutant genes, such as PSEN1, PSEN2, and APP, in iPSC-derived neurons provide exceptional platforms for elucidating disease mechanisms and formulating novel therapeutic strategies. Studies demonstrated that gene editing using CRISPR–Cas9 can mitigate the impairments linked to AD by addressing synaptic dysfunction and altered signaling pathways (108–111). Researchers have created gene-edited induced pluripotent stem cell models to investigate genetic mutations associated with AD, as well as knockout genes implicated in Alzheimer's pathogenesis, including ATP-binding cassette subfamily A member 7 (ABCA7) (112–114). The integration of animal models and CRISPR-edited iPSCs enhances scientists' understanding of AD risk factors and therapeutic options (115). CRISPR-engineered iPSC and organoid models have elucidated critical mechanisms in AD, including lipid metabolism (APOE), endosomal dysfunction (APP/PSEN1), electrophysiological abnormalities (PSEN2), and neuroimmune interactions (APOE Christchurch). These human-relevant systems are highly effective for identifying disease pathways and evaluating treatment alternatives (116–119). Stem cell models integrated with gene editing technologies facilitate the development of patient-specific therapies, paving the way for clinical applications.

Stem cell-derived models, in conjunction with gene editing, exhibit potential to enhance the development of tailored AD therapies. Table 2 Illiterates applications of CRISPR–Cas in AD. Utilizing CRISPR, scientists acquire enhanced insights into AD progression and develop successful therapeutic approaches.

Jia et al. (120) created a CRISPR-powered aptasensor that detects Aβ biomarkers via fluorescence detection via CRISPR-Cas12a. There is currently little study on the use of CRISPR-Cas13 and CRISPR-Cas3 in AD diagnosis. Although its potential in diagnostic domains has not been thoroughly investigated, Ca3′s primary DNA deletion function remains its most prominent feature.

PD, the second prevalent neurodegenerative ailment, is marked by the degradation of dopaminergic neurons, which results in particular motor symptoms such as rigidity, bradykinesia, and tremor (121). After the loss of neuronal, the Lewy bodies appeared with cytoplasmic inclusions, which are mostly composed of aggregates of α-synuclein protein misfolded and perhaps diffusion in a prion-like shape among interconnected areas synoptically (122). PD can be classified into two forms: sporadic and familial. The familial form accounts for only 10%−15% of all PD cases. The hereditary versions are attributed to mutations in multiple genes, including SNCA, parkin (PRKN), PTEN-induced kinase 1 (PINK1), DJ-1/Parkinson's disease protein 7 (PARK7), and LRRK2. SNCA and LRRK2 mutations cause autosomal-dominant types of PD, while PRKN, PINK1, and DJ-1 mutations cause autosomal recessive variants (123). Although the origins of sporadic forms are currently unknown, certain genetic and environmental risk factors have been discovered. For instance, genetic polymorphisms in the LRRK2 and SNCA genes elevate the likelihood of developing sporadic PD (124).

Between 1992 and 2021, global PD cases escalated from 450,000 to 1.34 million, with crude incidence rates increasing from 8.19 to 16.92 per 100,000 and age-standardized incidence rates rising from 11.54 to 15.63 per 100,000. Incidence rates increased significantly in persons aged 60 and beyond, reaching a zenith in those aged 85 and above. Forecasts indicate that by 2030, worldwide PD cases will total 1.93 million, with an age-standardized incidence rate of 27 per 100,000 (125).

PD is a ND mostly attributed to genetic abnormalities and environmental stresses (126). Genetic alterations in critical genes such as SNCA, LRRK2, PARK2, PINK1, and Glucocerebrosidase 1 (GBA1) are central to the disorder. Mutations in SNCA result in the improper folding of SNCA protein, leading to aggregation into Lewy bodies, which are indicative of PD in the brain (127). Mutations in LRRK2 that enhance gene function result in increased kinase activity, which is detrimental to neurons. Mutations in PARK2 and PINK1 disrupt mitochondrial quality control, leading to mitochondrial malfunction, poor mitophagy, oxidative stress, and ultimately, the demise of dopaminergic neurons (128). Neurons exhibit increased toxicity when mutations in the LRRK2 gene lead to the enzyme acquiring excessive kinase activity. The PARK2 gene regulates parkin activity, which acts as an E3 ubiquitin ligase; however, its dysfunction results in the buildup of protein damage (128). Neuronal damage exacerbates when GBA1 mutations induce mitochondrial malfunction and elevate oxidative stress inside cells (129, 130). Genetic abnormalities alter protein degradation pathways, impair mitochondrial function, and interfere with oxidative stress responses, ultimately leading to PD symptoms through the degeneration of dopaminergic neurons in the substantia nigra (131). Impaired mitochondria result in heightened oxidative stress and diminished ATP synthesis, causing neuronal damage. The development of PD occurs due to mutations in PINK1 and PRKN that impair mitochondrial quality control mechanisms (131). PD neuroinflammation occurs when activated microglia secrete tumor necrosis factor-alpha (TNF-α), interleukin (IL)-1β, and reactive oxygen species, which concurrently harm dopaminergic neurons. The stimulation of microglial cells by misfolded SNCA protein initiates a persistent inflammatory response. Extracellular immune cells, in conjunction with activated astrocytes, facilitate the progression of PD (132).

CRISPR–Cas techniques have proven essential in elucidating PD pathophysiology by facilitating the precise alteration of genes associated with dopaminergic neuron susceptibility (133). CRISPR-mediated knock-in and knock-out mice of LRRK2, PINK1, and PARK have clarified abnormalities in mitochondrial quality control and dysregulation of autophagy (134, 135). CRISPR correction of LRRK2-G2019S in patient-derived iPSCs reinstates dopaminergic differentiation and mitochondrial function, underscoring its therapeutic significance. CRISPRi has been utilized to diminish SNCA overexpression, hence decreasing aggregation and related toxicity (136). Novel therapeutic approaches encompass AAV-mediated CRISPR–Cas systems aimed at SNCA regulatory elements and base editors intended to rectify pathogenic point mutations; nonetheless, challenges related to delivery efficiency and off-target effects persist as significant obstacles (134).

CRISPR–Cas9 technologies, encompassing knockout, base editing, and gene correction, facilitate our comprehension of biological mechanisms and preserve phenotypes in models pertinent to PD. They possess significant potential for future gene therapy as they target genetic factors such as LRRK2, PINK1, and PARKIN, thereby aiding in the reduction of neuroinflammation and neuronal degeneration (137–140).

Recently, CRISPR–Cas9 genome editing has been employed in a laboratory to rectify mutations responsible for PD and to elucidate their pathogenic mechanisms. For example, employing CRISPR–Cas9 to eliminate the A53T-SNCA mutation resulted in a significant reduction in both SNCA mRNA expression and its protein levels (137, 141). Beligni et al. (142) demonstrated that CRISPR–Cas9 could rectify the LRRK2 p.Gly2019Ser (G2019S) mutation in patient-derived cells. This indicates its potential as a therapeutic agent to restore normal kinase function (143). A separate work employed CRISPR–Cas9 to entirely eliminate PINK1 in THP-1 cells, enhancing our understanding of PIN1′s function in maintaining mitochondrial integrity and the progression of PD (142). A recent work applied by (142) addresses the role of PINK1 in PD by employing CRISPR–Cas9 to generate PINK1 knock-out THP-1 cells. The PINK1 knockout exhibited diminished mitochondrial complex I activity alongside alterations in Tau protein expression patterns. In PINK1 knock-out macrophages, pro-inflammatory cytokines such as IL-6, IL-1β, and IL-23 were raised, while the lipid droplet count per cell decreased, indicating mitochondrial dysfunction and immune system pathological alterations (142).

Tokuda et al. expanded upon these findings by employing HDR to correct LRRK2 G2019S and truncate the LRRK2 kinase domain in marmoset embryonic stem cells and iPSCs. This rendered non-human primate cellular models applicable (144). In pertinent research, elevating glial cell line-derived neurotrophic factor (GDNF) levels in MPTP-treated neuronal cells via CRISPR–Cas9 was associated with reduced monoamine oxidase B (MAO-B) activity, diminished production of reactive oxygen species, and enhanced cell survival under neurotoxic stress (145). The in vitro applications of CRISPR–Cas9, including gene deletion, point mutation correction, gene knockout, and gene upregulation, demonstrate its versatility and efficacy as a tool for investigating the biology of PD and developing gene-based therapies.

Researchers have employed CRISPR–Cas9 to develop large-animal models (in vivo) of PD by introducing deleterious SNCA mutations into existing brain systems (146). Zhu et al. (146) employed CRISPR–Cas9 and SCNT to generate Guangxi Bama minipigs with the PD-associated E46K, H50Q, and G51D mutations in SNCA. Despite the presence of mutant SNCA in the early animals, they exhibited neither dopaminergic neurodegeneration nor Lewy body pathology at 3 months, highlighting the potential and limitations of this large-animal paradigm. Employing dCas9, CRISPRi effectively reduced SNCA mRNA and its protein levels in animal models (147). PINK1 and PARKIN constitute a pivotal signaling axis that significantly regulates the mitochondrial autophagy process in dopaminergic neurons. In PINK1 null Drosophila, the silencing of the ubiquitin c-terminal hydrolase L1 (UCHL1) gene using RNA interference can ameliorate the pathology associated with PD. The absence of UCH deubiquitination enhances mitochondrial autophagy by stimulating the expression of AMP-activated protein kinase (AMPK) and Unc-51–like autophagy-activating kinase 1 (ULK1) (148).

Researchers have developed the mitochondrial complex I (MCI)-Park model in mice using CRISPR Cas9 to generate dopamine neurons deficient in NADH dehydrogenase [ubiquinone] iron–sulfur protein 2 (NDUSF2), which encodes mitochondrial complex I. Mice deficient in NDUSF2 exhibited neurodegenerative alterations (149). Point mutations in the vacuolar protein sorting 35 gene (VPS35) are linked to autosomal dominant late-onset PD (PARK17). The homozygous deletion of Vps35 using CRISPR Cas9 led to a survival disadvantage and a marked decrease in dopamine release in the caudate putamen of adult homozygous Vps35 mutant mice (150). Peroxidized phospholipids resulting from ferroptosis have been linked to the beginning of certain PD. Patatin-like phospholipase domain-containing protein 9 (PNPLA9), functioning as a hydrolytic enzyme, exhibits a preference for hydrolyzing peroxidized phospholipids. In mice, the ablation of PNPLA9 led to the advancement of Parkinsonian motor abnormalities and the accumulation of peroxidized phospholipids (151). This represents a promising method for gene repression in vivo PD models.

Researchers have investigated the application of CRISPR techniques to activate genes. Narváez-Pérez et al. employed a pseudo-lentivirus to deliver a CRISPR/SAM system to rat astrocytes, enhancing the expression of tyrosine hydroxylase and resulting in dopamine production. When these altered astrocytes were introduced into a rat model exhibiting 6-hydroxydopamine (6-OHDA)-induced injury, they improved motor function (138). This demonstrated an innovative approach to treating PD. Increased CRISPR-Cas9 genome editing has produced mouse models that eliminate NDUSF2 (mitochondrial complex I) or knock out VPS35 and Pnpla9 (148–152). This induces neurodegeneration and motor dysfunction akin to the manifestations of PD. Conversely, the ablation of PRKN, PINK1, and DJ-1 thrice in murine models did not induce neurodegeneration, indicating that some species exhibit greater resilience and that the genetic modeling of PD is complex (153).

iPSC models derived from individuals with PD are valuable for investigating genetic variations associated with the disorder, such as LRRK2, SNCA, PARKIN, and ubiquinol–cytochrome c reductase core protein 1 (UQCRC1) (154–156). Neural stem cells (NSCs) harboring the LRRK2 p.G2019S mutation are predisposed to proteasome stress, nuclear envelope dysfunction, clonal proliferation issues, and challenges in neural differentiation (157).

Moreover, rectifying this mutation using knock-in techniques restores the phenotype to normal, indicating that nuclear morphology may serve as a biomarker for PD. LRRK2 p.G2019S disrupts mitochondrial mitophagy by impairing the Miro–PINK1/PARKIN pathway, hence hindering cellular respiration and metabolism (158). Similarly, NSCs derived from individuals with PARK2 mutations differentiate into neurons exhibiting elevated oxidative stress, activated Nrf2 pathways, and abnormal mitochondrial morphology and dysregulated homeostasis (159, 160).

Research on dopaminergic SH-SY5Y cells with UQCRC1 mutations, observed in both familial and sporadic PD, indicates that axonal degradation occurs and mitochondrial respiratory chain function ceases. These iPSC-derived cellular models demonstrate how genetic risk factors influence pathways governing proteostasis, mitochondrial integrity, oxidative equilibrium, and cellular morphology (161). They demonstrate the utility of CRISPR–Cas9-mediated gene repair or knockout in restoring cellular function, thereby enhancing the understanding of underlying mechanisms and identifying potential therapeutic targets in PD.

CRISPR-modified PD organoids replicate dopaminergic neuron degeneration, oxidative stress, and α-synuclein pathology, offering a three-dimensional framework for longitudinal investigations. Three-dimensional human midbrain organoids generated from CRISPR-modified induced pluripotent stem cells augment the physiological relevance of these models. These organoids have a spatial organization akin to the human substantia nigra, with dopamine-producing neurons, astrocytes, and nascent microglial-like populations. CRISPR-induced PD mutations in organoids expedite SNCA fibrillization, impair neuromelanin production, cause lysosomal dysfunction, and lead to progressive neuronal loss, reflecting clinical illness with temporal advancement. These 3D systems enable longitudinal examination of synapse degeneration, neuroinflammation, and intercellular communication processes inadequately represented in 2D cultures (162).

Biomedical applications illustrate the usefulness of CRISPR-Cas12 technology, which has shown promise in research for treating NDs, including PD. Healthcare research continues to develop and still under research, studies on the applicability of CRISPR-Cas12 for PD treatment, utilizing the existing CRISPR technology still limited.

Researchers have discovered intriguing applications for CRISPR-Cas3′s RNA-targeting DNA editing technologies in treating NDs, including PD. CRISPR systems based on DNA editing fail to target single-stranded RNA, but Cas13 can precisely control gene expression using RNA targets that do not alter genomic sequences (163). The application on PD and CRISPR-Cas13 still limited and shown in Table 3.

HD is a progressive ND, an autosomal dominant disease. The condition is attributed to an enlarged cytosine-guanine-adenine (CAG) repeat in the HTT gene located on chromosome 4 (4p16.3), resulting in a mutant huntingtin (mHTT) protein characterized by an elongated polyglutamine tract (164–166). The deleterious gain-of-function characteristics of mHTT initiate a series of molecular disruptions that ultimately result in the specific destruction of medium spiny neurons within the striatum. At the molecular level, mHTT impairs transcriptional control by anomalously interacting with transcription factors, including CREB-binding protein (CBP), resulting in extensive transcriptional suppression and modified neuronal identity. Moreover, mHTT disrupts axonal transport by obstructing microtubule-associated motor proteins (dynein/dynactin), leading to impaired trafficking of vesicles, mitochondria, and neurotrophic signals such as brain-derived neurotrophic factor (BDNF) (167). Individuals with optimal health typically possess 16–20 CAG repeats, however, those afflicted with HD may have over 40 repetitions. Extended repeats are associated with earlier onset and more severe symptoms (164). The misfolded mHTT protein forms toxic aggregates that disrupt critical cellular activities such as transcription, energy metabolism, axonal transport, and apoptosis. This ultimately results in the demise of striatal and cortical neurons. In the clinic, HD manifests in middle age with issues related to movement, cognition, and mental health (168). The studies detected the mitochondrial dysfunction is a primary characteristic, marked by diminished respiratory chain activity, compromised calcium buffering, elevated oxidative stress, and increased vulnerability to apoptosis. mHTT also undermines autophagy and proteostasis by obstructing autophagosome cargo loading, impairing ubiquitin–proteasome functionality, and facilitating the formation of insoluble aggregates. Synaptic dysfunction manifests early via altered NMDA receptor activation, excitotoxicity, and compromised corticostriatal connection. Recent data indicates that non-cell-autonomous mechanisms, including as impaired astrocytic glutamate absorption and microglial activation, contribute to the acceleration of neuronal death. These interrelated pathways collectively characterize HD as a progressive multisystem condition influenced by proteotoxicity, transcriptional dysregulation, compromised energy metabolism, and synaptic susceptibility (169, 170). A 2012 meta-analysis revealed significant regional variation in the prevalence of HD, with elevated rates in Europe, North America, and Australia (5.7 per 100,000) compared to diminished rates in Asian populations (0.4 per 100,000) (171).

HD pathophysiology comes from the misfolding of mHTT into aggregates that impair cellular function, ultimately leading to neuronal cell death (172). The protein interferes with transcriptional regulators, leading to aberrant gene expression throughout the organism (173). Mitochondrial dysfunction, along with elevated oxidative stress, induces neuronal injury by diminishing cellular energy levels (174). Concurrent aberrant glutamate activity induces neuronal death pathways. The mHTT protein induces malfunction in the proteasome and autophagy processes, leading to the accumulation of toxic proteins (175). Neuronal damage increases due to chronic neuroinflammation caused by activated microglia and astrocytes in the nervous system (176). The prospective applications of CRISPR-Cas technology for the treatment of HD are significant due to its capacity for precise genetic material alteration. Gene therapy employing targeted CRISPR-Cas9 serves as a prospective treatment strategy for disease management by directly correcting genetic mutations at their origin (164, 177). Various studies have been demonstrated the potential of CRISR/Cas tool in HD application and theoretical aspects such as gene editing, selective targeting of mutant alleles, and excision of expanded cag repeats (178–180). Shin et al. undertook a study with the goal of enhancing the specificity of alleles. They employed a customized approach using CRISPR–Cas9 that selectively targeted specific alleles by modifying PAM-altering single nucleotide polymorphism (SNP). This approach focused on identifying CRISPR–Cas9 sites that were unique to each patient and utilized a comprehensive understanding of the haplotype structure of the HTT gene. The objective was to deactivate the mutated HTT allele for a specific haplotype specifically (181).

Researchers have employed CRISPR–Cas9 genome editing to create isogenic cell models (in vitro) of HD by precisely modifying the CAG repeat region in the HTT gene. CRISPR has been used to allele-specific targeting and genetic correction, Allele-specific excision of the mutant HTT, Shin employed SNP-guided dual-CRISPR–Cas9 targeting to excise approximately 44 kb from the mHTT allele in patient-derived cells. This eliminated mutant mRNA and protein while preserving the wild-type allele (181). In a separate investigation, Yang et al. showed that the application of CRISPR–Cas9 in HD140Q-KI mice to remove HTT in a non-allele-specific way can successfully and permanently eradicate the harmful effects of polyglutamine expansion on neurons in the adult brain. The study population showed a significant reduction in reactive astrocytes and a notable improvement in motor impairment (182). Another study utilized CRISPR to CAG-repeat excision, such as a study introduced 150 CAG repeats into exon 1 of HEK293T cells to create clones resembling the condition, perhaps facilitating the investigation of disease mechanisms (183). In pertinent research, scholars employed Cas9 ribonucleoprotein complexes and single-stranded oligodeoxynucleotide donors to create HEK293T lines containing 41, 53, and 84 CAG repeats. This facilitated the examination of alterations in cellular phenotypes associated with varying repeat lengths and contributed to the development of novel treatments (184). Another approach employed Cas9 nickases and guide RNAs to excise CAG repeats, resulting in frameshifts that activated premature stop codons and inhibited mHTT production. This illustrates how accurate genome editing could reinstate the typical allele configuration (185). In transgenic BacHD mice, Monteys et al. (186) incorporated a modified human HD allele and targeted human mHTT exon 1. BacHD animals treated with rAAV.spCas9 and rAAV.sgHD1B/i3 showed decreased HTT mRNA levels in their right hemisphere but not in their uninfected left (186). Ekman et al. (187) revealed effectively inhibited the expression of the human mutant HTT gene in R6/2 mice that carried human mHTT exon 1 with roughly 115–150 CAG repeats in 2019 using AAV-mediated CRISPR-Cas9. This study showed that R6/2 mice's lifespan and motor deficits were enhanced by inhibiting HTT expression. In conclusion, it has been shown in various animal models that HD may be effectively treated by employing CRISPR-Cas9 to eliminate mHTT gene expression (187).

CRISPR-based gene-silencing techniques have been employed to reduce the expression of mHTT with the direct deletion of repeat expansions. CRISPRi exclusively inhibited mHTT transcription in fibroblasts derived from HD patients, without affecting wild-type HTT (188). Previous work employed CasRx (RNA-targeting Cas13) to downregulate HTT exon 1 in HEK293T cells, HD 140Q knock-in mice, and HD knock-in pigs. This decreased mHTT levels, diminished gliosis, and decelerated neurodegeneration (189). Ekman et al. (187) revealed effectively inhibited the expression of the human mHTT gene in R6/2 mice that carried human mHTT exon 1 with roughly 115–150 CAG repeats in 2019 using AAV-mediated CRISPR-Cas9. This study showed that R6/2 mice's lifespan and motor deficits were enhanced by inhibiting HTT expression. It has been shown in various animal models that HD may be effectively treated by employing CRISPR-Cas9 to eliminate mHTT gene expression (187).

Oura et al. (190) addressed the target range limits of WT-SpCas9, scientists engineered SpCas9-NG, which detects NGN PAMs. They used SpCas9-NG to precisely contract HD CAG repeat tracts in mouse embryonic stem cells. The procedure of rectifying mutations restored distinctive characteristics as well as health conditions in both neurons derived from modified stem cells and the animals produced by these cells. This study reveals that SpCas9-NG is a powerful way to treating enlarged CAG repeats and other disease variants that are difficult to target (190). Another study via Merienne et al. (191) found the development of the KamiCas9 self-inactivating CRISPR–Cas9 system, which enables high-efficiency Cas9 expression during transitory intervals. Using their technique on adult mouse neuronal and glial cells, researchers were able to inactivate the HTT gene while concurrently improving HD disease markers. The KamiCas9 system made it possible to reduce unwanted edits when evaluating human iPSC-derived cells since it had less off-target effects than typical constitutive Cas9 systems. The study shows how self-inactivating CRISPR–Cas9 technologies could become a feasible therapy option for NDs (191). Ultimately, allele-specific CRISPR–Cas9 editing utilizing SNP-generated PAM sites facilitated the targeted silencing of mHTT by nonsense-mediated degradation in approximately 20% of European HD patients. This demonstrated exceptional accuracy and little off-target effects following GUIDE-Seq validation. This research demonstrates the versatility of CRISPR approaches in investigating the etiology of HD and formulating novel therapies. They can be utilized for genome editing, gene silencing, and allele-specific strategies (192).

CRISPR-based techniques have been shown to be effective in vivo for HD by reducing levels of mHTT in the brain. Bunting and Donaldson demonstrated that employing antisense oligonucleotides to inhibit MSH3 activity reduced HTT CAG repeat expansions in HD mouse models. This indicates that gene-targeting methodologies may alter the trajectory of the disease (193). Similarly, administering CRISPR–Cas9 to striatal neurons in HD140Q-knockin mice significantly reduced endogenous mHTT, inhibited aggregation formation, and ameliorated early symptoms of neurodegeneration and motor dysfunction, all without causing neuronal damage (182). In the prevalent R6/2 mouse model, CRISPR–Cas9-induced deletion of HTT resulted in reduced protein inclusions, enhanced motor function, and an extended longevity. This indicates that targeted genome editing may be an effective approach for treating HD (103, 186, 187, 191).

Enhanced editing precision has significantly improved safety and efficacy. Base editing was utilized in vivo to disrupt caspase-6 cleavage sites within the HTT gene. This resulted in mHTT isoforms that exhibit resistance to proteolytic processing. This approach effectively reduced the detrimental aggregation of huntingtin and safeguarded against neuronal loss in the striatum and brain of HD animal models (187). In vivo investigations, encompassing antisense, CRISPR–Cas9 gene disruption, and base editing, demonstrate that reducing or altering mHTT expression can decelerate the advancement of neuropathological disorders and enhance functional outcomes. This indicates that CRISPR possesses the potential to serve as a disease-modifying instrument for HD (194).

The integration of iPSCs with CRISPR–Cas technology has advanced HD research by creating precise, isogenic models and revitalizing cells. Scientists developed homozygous HEK293T cell lines with varying amounts of CAG repeats in the HTT gene to facilitate mechanistic investigations and drug screening (184). Researchers have employed CRISPR–Cas9 to rectify pathogenic CAG expansions in iPSCs, effectively reversing initial indicators of mitochondrial dysfunction in HD-derived neural stem cells, including respiratory issues, elevated reactive oxygen species, and calcium dysregulation (195–198). The results indicate that the selective elimination of CAG repeats can restore mitochondrial function and maintain cellular equilibrium. CRISPR–Cas9 editing facilitates allele-specific disruption, repeat excision, and gene repair in both in vitro and in vivo environments. Outstanding issues remain, including its efficacy, long-term safety, and performance on alternative genes. This gene-editing technique possesses significant potential to alter the trajectory of HD (199). Table 4 illustrates some HD applications and consequences.

Recent advancements indicate that CRISPR-Cas13 systems may be effective in treating HD by targeting RNA transcripts associated with the disorder. Morelli and Wu (152) conducted a notable study that advanced an enhanced CRISPR-Cas13d system known as Cas13d-CAGEX. This treatment led to enduring enhancements in motor function, reduced neuronal degeneration, and sustained therapeutic effects with minimal adverse effects observed for up to 8 months post-injection (200).

The efficient application of CRISPR-Cas13d demonstrates its considerable potential as an RNA-targeting tool for HD and possibly other dominant genetic disorders. Cas13d represents a safer alternative for prolonged gene regulation as it specifically inhibits pathogenic transcripts without altering genomic sequences. This differs from DNA-targeting techniques. Numerous studies employ CRISPR-Cas13 in Huntington's disease, although few utilize CRISPR-Cas12. No studies have utilized CRISPR-Cas3 in HD research, indicating that these methods remain underexplored for neurodegenerative therapeutics.

ALS is a progressive ND characterized by the degeneration of both upper and lower motor neurons. This results in muscular weakening, atrophy, paralysis, and ultimately respiratory failure (201–203). Epidemiological studies indicate that incidence significantly varies by geography, with rates between 0.26 and 23.46 cases per 100,000 person-years, whereas prevalence ranges from 1.57 to 11.80 instances per 100,000 individuals across various countries (203, 204). Approximately 75% of ALS cases occur sporadically, although around 25% are familial and associated with over 135 genetic loci (205–208). A defining characteristic of ALS is the cytoplasmic mis localization and aggregation of TDP-43, which impairs RNA splicing, transport, and stability, resulting in extensive transcriptome dysregulation (202, 209). Mutant superoxide dismutase 1 (SOD1) and fused in sarcoma (FUS) intensify cellular stress through protein misfolding, oxidative injury, and impaired DNA damage repair. Intrinsic neuronal abnormalities are exacerbated by non-cell autonomous mechanisms, such as microglial activation, astrocytic glutamate toxicity due to diminished excitatory amino acid transporter 2 (EAAT2) expression, mitochondrial dysfunction, and compromised axonal transport. These mechanisms collectively begin a self-sustaining loop of neuroinflammation, energy depletion, and synaptic degeneration, ultimately resulting in motor neuron death (210, 211). The hexanucleotide (GGGGCC) expansion in C9orf72 is the predominant mutation among these. It induces toxic RNA foci and dipeptide repeat protein accumulation in around 40% of familial cases and 7% of sporadic cases (212–214). TAR DNA-binding protein (TARDBP) mutations disrupt TDP-43 protein homeostasis (4% familial, 1% sporadic) (215, 216), while SOD1 mutations lead to oxidase dysregulation and protein aggregation (217, 218) in 12% of familial cases and 1%−2% of sporadic cases. Moreover, alterations in FUS, heterogeneous nuclear ribonucleoprotein A1 (HNRNPA1), profilin 1 (PFN1), and kinesin family member 5A (KIF5A) influence RNA metabolism, axonal transport, and cytoskeletal architecture, ultimately resulting in motor neuron degeneration in ALS (219–223).

The CRISPR–Cas genome-editing technology offers a significant method for developing targeted ALS therapies through precise genetic modifications (224). The protentional applications of CRSPR/Cas in ALS disease was gene silencing and correction, disease modeling, gene therapy delivery (16, 224–226) (Figure 3). In vivo CRISPR techniques have been employed to directly target pathogenic ALS mutations in animal models, yielding quantifiable functional improvements and molecular restoration. AAV-mediated CRISPR–Cas9 targeting of mutant SOD1 decreased mutant SOD1 levels of proteins in the spinal cord and enhanced motor function and survival in transgenic mice, illustrating that somatic gene disruption can alleviate disease symptoms. Likewise, the excision of the pathogenic GGGGCC hexanucleotide repeat expansion (HRE) in C9orf72 by paired CRISPR–Cas9 guides diminished RNA foci and dipeptide repeat proteins, therefore ameliorating molecular abnormalities in various transgenic mice models (227). Recently, RNA-targeting CRISPR systems (Cas13/CasRx) designed for high fidelity have been utilized in vivo to diminish harmful repetitive RNAs and subsequent dipeptide repeat proteins (DPR) toxicity, providing a non-DNA-cleaving therapeutic approach that may mitigate irreversible on-target genetic concerns. These investigations demonstrate the feasibility of in vivo CRISPR interventions to reverse fundamental ALS processes, while also emphasizing the problems of delivery, immunogenicity, and product purity (editing byproducts) that persist for clinical application (227).

In vitro CRISPR modification of human cells, particularly patient-derived fibroblasts and iPSC-derived motor neurons, has proved essential for elucidating ALS pathophysiology and confirming targets. The CRISPR–Cas9-mediated removal or allelic modification of C9orf72 repetitions in patient-derived iPSC neurons diminishes RNA foci and DPR accumulation, while reinstating nucleocytoplasmic transport and cellular survival. CRISPR-induced knockouts of SOD1, FUS, or TARDBP in human cellular models replicate key mitochondrial, RNA metabolism, and proteostasis deficiencies, facilitating mechanistic comparisons among genotypes. The in vitro application of RNA-targeting Cas13 devices enables the direct reduction of hazardous repetitive RNAs in patient cells, facilitating swift assessments of guide efficacy and collateral activity prior to in vivo experimentation. Cell-based research is essential for determining dosage, assessing off-target effects, and developing biomarkers (228).

CRISPR–Cas9 technology has demonstrated its ability to rectify mutations associated with ALS in iPSC in vitro models derived from patient cells. For example, employing Cas9 along with a co-transfected donor template to rectify the SOD1 A272C mutation in fibroblast-derived iPSCs restored the wild-type sequence (229). Similarly, rectifying the SOD1 E100G mutation in iPSCs using homology-directed repair not only increased the quantity of motor neurons, the size of the soma, and the formation of neurites, but also diminished cell mortality post-differentiation (230). In C9orf72-associated ALS, employing CRISPR–CasRx to diminish sense and antisense repeat transcripts in HEK293T cells and patient-derived iPSC neurons decreased RNA foci and dipeptide repeat proteins, thereby providing protection against glutamate-induced excitotoxicity. Furthermore, utilizing Cas9 HDR to accurately excise the deleterious HRE from C9orf72 maintained the gene's integrity, restored expression and methylation levels to baseline, reduced intron retention, and eliminated detrimental repeat artifacts in motor neurons derived from iPSCs. All of these in vitro experiments demonstrate that CRISPR-based editing is a versatile and promising approach to address the primary genetic causes of ALS (231). The generation of SOD1+/A272C iPSCs and FUS+/G1566A iPSCs from primary fibroblasts of two ALS individuals, one possessing a heterozygous A272C mutation in the SOD1 gene and the other a heterozygous G1566A mutation in the FUS gene. The CRISPR-Cas9 approach effectively rectified the SOD1+/A272C and FUS+/G1566A mutations in ALS iPSCs utilizing a donated plasmid or single-stranded oligodeoxynucleotide (ssODN) as the repair template, resulting in restored ALS iPSCs that exhibited normal pluripotency. Furthermore, SOD1+/A272C and repaired iPSCs were prompted to differentiate into motor neurons, resulting in the identification of 899 aberrant transcripts using RNA sequencing. This work may assist in identifying biomarkers and pathways for ALS and provides novel evidence for ALS therapy (229).

Base editing and CRISPR–Cas9 are two efficacious techniques for investigating and addressing ALS, particularly in vivo animals. An essential approach involves utilizing AAV9 to administer CRISPR components to transgenic ALS mice with the gain-of-function SOD1 G93A mutation. Initial endeavors to employ S. aureus Cas9 (SaCas9) and gRNAs for enhancing survival and muscle preservation yielded limited success; hence, alternative strategies were explored. Base editors, including cytidine or adenine deaminase linked to a nickase Cas, enable accurate single-nucleotide modifications without generating double-strand breaks an advantageous characteristic for rectifying or deactivating point mutations in familial ALS. A pivotal proof-of-concept study using adenine base editing to inactivate a mutant SOD1 allele in vivo, enhancing motor performance in an ALS mouse model. This demonstrated that base editing can effectively diminish hazardous protein levels and yield phenotypic advantages while circumventing several consequences associated with DSBs. Base editing is limited by editable sequence contexts and bystander edits, while the delivery of base-editor cargo to motor neurons presents a translational bottleneck; yet, these results demonstrate substantial potential for therapy for point-mutation ALS. Researchers employed a dual AAV9 split-intein methodology to circumvent the limitations of vector size, enabling the creation of specific nonsense mutations in SOD1 (227, 232, 233). We must exercise caution in simultaneous initiatives aimed at targeting FUS and TARDBP mutations to avoid the detrimental consequences of entirely eliminating a gene (234, 235). Despite these challenges, employing CRISPR to reduce mutant SOD1 in G93A-SOD1 mice significantly improved their motor functions, postponed illness onset by 37%, and extended their longevity by 25%. This indicates that gene editing possesses therapeutic potential (227).

CRISPR–Cas9 has been effectively employed to rectify faulty expansions in the C9orf72 gene, the most common genetic contributor to ALS. This pertains to SOD1-related ALS. Multiple groups have utilized paired guide RNAs around the hexanucleotide repeats to induce excision by NHEJ in iPSCs derived from patients. They have successfully eliminated up to 11% of the target DNA. Unanticipated deletions occurred; yet, C9orf72 expression remained constant (236–238). To enhance fidelity, normal-length repeats were included by HDR techniques, restoring typical methylation and transcription profiles. Neurons derived from these modified iPSCs exhibited a reduced number of RNA foci, diminished dipeptide repeat protein aggregates, and enhanced stress resilience (231, 239). SOD1 participates in the detoxification of superoxide radicals, and mutations in SOD1 lead to enhanced harmful activity (209, 240). The G93A-SOD1 mouse model possesses roughly 25 tandem repeat copies of the human SOD1G93A transgene, demonstrating signs of amyotrophy and dyskinesia observed in ALS patients (241). CRISPR modification of iPSCs has emerged as a fundamental element in ALS research. By producing isogenic iPSC pairs (mutant vs. corrected control), researchers can associate genotype with phenotype while minimizing background interference. The differentiation into motor neurons and three-dimensional spinal/brain organoids facilitates the replication of TDP-43 mislocalization, RNA foci, DPR synthesis, mitochondrial dysfunction, axonal transport abnormalities, and non-cell-autonomous contributions from astrocytes and microglia found in ALS. These CRISPR-iPSC platforms are extensively utilized for mechanistic analysis, high-throughput drug screening, and preclinical assessment of gene-editing methodologies, including the evaluation of AAV/SNP-guided allele selectivity, base editors, and Cas13 guides. iPSC platforms are essential for precise off-target and transcriptome-wide RNA-editing evaluations before animal or clinical use (226).

Recent advancements in CRISPR-based RNA-targeting methodologies suggest their potential utility in the treatment of ALS. Zhang et al. (2021) in their review article discussed an enhanced variant of CRISPR-Cas13d, termed CasRx, which can degrade both sense and antisense C9ORF72 repeat RNAs while preserving normal gene expression. Utilizing AAV vectors, CasRx was administered to HEK cells and patient-derived iPSC neurons in vitro, as well as to transgenic animal models in vivo. This resulted in significant reductions in RNA foci, DPRs, and glutamate-induced excitotoxicity (163). In another study, researchers employed Cas13 and Cas7-11 effectors to target ataxin-2 in models of TDP-43 proteinopathy. Direct injection of the Cas13 system into mice effectively reduced TDP-43 aggregation and the location of stress granules. This resulted in improved behavior, increased longevity, and reduced neurodegeneration in the mice. In comparison to previous Cas7-11 tools (242), Cas13 variants exhibiting enhanced fidelity demonstrated superior transcriptome specificity. The results indicate that RNA-targeting CRISPR systems possess significant potential as a therapeutic approach for ALS, as they can target critical disease characteristics. Table 5 illustrates CRISPR techniques in ALS.

The efficient and selective delivery to certain cell types is the primary practical limitation for CNS-targeted CRISPR therapies. AAV vectors provide strong neuronal tropism, sustained transgene expression, and established manufacturing processes, rendering them the primary tool for in vivo CNS editing; however, their restricted packaging capacity hinders the delivery of larger editors (such as prime editors and certain base-editor constructs), and continuous Cas expression from AAV may elevate off-target activity and immunogenicity (243). Lipid nanoparticles (LNPs) offer a supplementary, transient-expression approach for delivering mRNA or RNPs, thereby minimizing the editor's exposure duration. Recent advancements in LNP formulations have facilitated systemic and targeted delivery in preclinical models; however, efficient BBB penetration and neuronal specificity continue to be areas requiring further optimization (244). Engineered extracellular vesicles/exosomes represent a novel option characterized by low immunogenicity and potential for targeted cell delivery; ongoing research emphasizes scalable loading techniques, surface modification for tropism, and comprehensive biodistribution characterization (245). A pragmatic hybrid approach such as compact editors encapsulated in AAV for localized distribution, LNPs for temporary systemic administration, or exosome-based vehicles.

Long-term safety issues associated with CRISPR encompass not just off-target mutations but also significant on-target deletions, insertions, inversions, chromosomal translocations, and other intricate structural changes that conventional PCR techniques may overlook. Identifying these occurrences necessitates genome-wide, high-resolution methodologies, including long-read sequencing and capture-based assays. Strategies for risk mitigation encompass the temporary administration of CRISPR components (RNP or mRNA), the utilization of high-fidelity or double-strand break-free editors (nickase, base, or prime editors), and meticulous optimization of guide RNA, bolstered by empirical off-target analysis. Comprehensive preclinical genotoxicity assessment and prolonged molecular and clinical surveillance, with established cessation criteria, are crucial for initial human studies (268).

Both the adaptive and innate immune responses present significant challenges for repeated or systemic CRISPR therapy. Documented pre-existing humoral and cellular immunity to AAV capsids and frequently utilized Cas orthologs (SpCas9, SaCas9) can neutralize vectors or induce T-cell-mediated clearance of edited cells; additionally, sustained expression of bacterial nucleases extends the period for immune recognition (293). Effective mitigation strategies, substantiated by preclinical and early clinical studies, encompass the utilization of less common AAV serotypes or engineered capsids, transient immunosuppression during dosing, the administration of editors as mRNA or RNPs (via LNPs or alternative transient carriers) to minimize antigen exposure, and the creation of Cas variants featuring diminished immunogenic epitopes or humanized sequences. Thorough immune profiling in pertinent animal models and patient samples is essential for formulating appropriate therapeutic regimens (294).

Importantly, although pre-existing humoral and cellular immunity to commonly used Cas orthologs (e.g., SpCas9/SaCas9) has been documented in human populations (295), published clinical experience with CRISPR–Cas9 therapies to date has not reported clinically meaningful Cas9-directed immune toxicities (296). For example, in the first systemic in vivo CRISPR–Cas9 trials targeting transthyretin (TTR) (LPN delivery of Cas9 mRNA and sgRNA; NCT04601051), reported treatment-related events were primarily transient infusion-related reactions (and transient liver-enzyme elevations in some participants), without persistent immune-mediated sequelae attributed specifically to Cas9 (297). In parallel, ex vivo CRISPR–Cas9 edited autologous hematopoietic stem and progenitor cells products (e.g., Exagamglogene autotemcel) have shown safety profiles largely consistent with conditioning/transplant procedures rather than editor immunogenicity, which is biologically plausible given the transient exposure to Cas9 during cell manufacturing (298).

Nevertheless, ongoing and future trials, particularly those requiring CNS delivery and/or potential redosing, should continue systematic immune surveillance (anti-Cas antibodies, T-cell responses, cytokine profiles) to define risk more precisely in neurological indications.

CRISPR-based therapeutics for NDs presents significant ethical and regulatory issues, especially when utilized solely on somatic cells. Critical concerns encompass securing informed consent for permanent genomic modifications, equitable access to expensive therapies, and meticulously assessing the risks and benefits of irreversible operations in the CNS. First-in-human trials necessitate stringent supervision, particularly when employing novel delivery technologies. Despite widespread rejection of germline editing, its public discourse continues to influence trust in somatic editing. Robust community involvement, autonomous ethical assessment, and compliance with international standards are needed. Regulators are progressively mandating comprehensive preclinical evidence about safety, immunological responses, and off-target effects, in addition to dependable manufacturing and long-term monitoring strategies. Engaging bioethicists and patient representatives in trial design enhances the quality of consent and public acceptance (320).

Although CRISPR–Cas possesses considerable promise for the treatment of various genetic disorders, issues concerning off-target effects and the efficient delivery of the CRISPR–Cas genome editing apparatus must be resolved to facilitate its implementation in clinical settings, particularly for neurodegenerative diseases. The primary problem in creating theragnostic for brain injury is site-specific delivery. Instances have arisen throughout the years in which the transport of nanoparticles (CRISPR–Gold, magneto-electric nanoparticles) has been effectively utilized (303, 364). This technique facilitates genomic modifications, encompassing the deletion of extensive nucleotide sequences, homologous recombination, insertion/deletion point mutations, and transcriptional regulation of certain genetic regions. Owing to its functional characteristics and resilience to epigenetic alterations, CRISPR–Cas9 has emerged as the most advantageous method for genome editing in human disease models, both in vivo and in vitro (365). Nonetheless, as previously discussion, the primary barrier of effective gene therapy for neurodegenerative diseases is the inadequate comprehension of its pathogenic mechanisms. Nonetheless, with a growing quantity of research aimed at elucidating the molecular causes and formulating gene therapies for this disease indicates that CRISPR–Cas9 technology has considerable promise for enhancing our comprehension of NDs and for effective future interventions targeting specific genes.

Future applications of CRISPR–Cas focus on precise, allele-specific editing to selectively target mutant alleles responsible for autosomal dominant NDs such as HD and fAD. Advancements in base editors and prime editing facilitate single-nucleotide modifications without inducing double-stranded breaks, hence reducing off-target effects and enhancing safety profiles (54). In preclinical models of HD, the disruption of the mHTT gene with CRISPR–Cas9 has shown promising results, including improved motor performance and reduced neurotoxicity (186, 366).

Recent preclinical studies indicate that genome- and transcriptome-targeted CRISPR approaches can significantly alter ALS biology in vivo and in human cells. Specifically, in vivo base-editing or nuclease techniques that reduce mutant SOD1 expression have demonstrated functional advantages and prolonged survival in murine models, thereby providing robust therapeutic proof-of-concept for gene-specific familial ALS (233). Similarly, the excision of the C9ORF72 GGGGCC repeat expansion via Cas9 diminishes RNA foci and DPRs in cellular and murine models, whereas RNA-targeting CRISPR (Cas13/CasRx) can reduce toxic repeat RNAs/DPRs without modifying DNA providing a complementary, less-permanent approach for patients (228).

Implementing these advancements in clinical settings will necessitate concurrent advancements in four domains. The primary obstacle is the secure and effective delivery to motor neurons (spinal cord and pertinent brain regions) at a therapeutic scale: innovative AAV capsids and non-viral methods (LNPs, nanoparticles, intrathecal or intraparenchymal regional techniques) must be refined for biodistribution, dosage, and immunogenicity in large animals prior to human trials (367). Secondly, it is imperative to minimize genotoxic risk by incorporating base and prime editors that circumvent double-strand breaks, utilizing transient delivery formats (such as RNPs, self-inactivating vectors, and split-intein systems), and conducting thorough, unbiased genome- and transcriptome-wide off-target profiling within preclinical packages (233).

Third, the clinical translation is enhanced by established regulatory and biomarker precedents: the approval and regulatory experience with antisense oligonucleotide therapy for SOD1-ALS (tofersen) illustrates that reducing a disease protein and exhibiting biomarker effects (e.g., neurofilament light) can facilitate accelerated pathways while long-term clinical benefits are validated in confirmatory studies. This presents three practical implications for CRISPR strategies: (a) initial trials should incorporate robust molecular biomarkers (Cerebrospinal fluid (CSF)/plasma neurofilament and target RNA/protein measurements), (b) patient selection must closely correspond with the genomic target (e.g., SOD1 or chromosome 9 open reading frame 72 (C9ORF72) carriers), and (c) early regulatory engagement in development is essential (368).

One of the biggest challenges still is effective and focused in vivo distribution of CRISPR components. Developed to improve transport over the BBB to reach particular neural populations are innovations include AAV vectors, LNPs, and modified exosomes (224, 369–371). To fix mutations in SOD1 in ALS mice, for instance, a dual AAV system has been employed to deliver CRISPR–Cas9, hence displaying longer longevity (232). Furthermore, in development are ligand-conjugated nanoparticles to increase CNS cell-type selectivity (372).

CRISPR/dCas9, in conjunction with transcriptional repressors or activators, facilitates epigenetic modulation of gene expression without cleaving DNA. This approach offers a reversible and perhaps safer alternative, especially for multifactorial conditions like PD or sporadic AD. Research has demonstrated that in PD models, dCas9-mediated activation of neuroprotective genes (e.g., BDNF) can preserve dopaminergic neurons (373).

The advancement of human-relevant disease models is being revolutionized by CRISPR–Cas technology. Modifying disease-related mutations in iPSCs enables them to emulate the pathogenic characteristics of neurodegeneration. Novel therapeutic targets and disease development regulators are being identified using genome-wide CRISPR screening (374). These tools facilitate the identification of significant genetic associations and cellular pathways involved in disease mechanisms.

The integration of CRISPR-based methodologies with other treatment strategies such as RNA interference (RNAi), antisense oligonucleotides (ASOs), and small molecule pharmaceuticals is a significant trend. This combination may mitigate CRISPR-induced risks while simultaneously enhancing therapeutic efficacy. In AD models, targeting both APP and BACE1 genes has demonstrated a synergistic reduction in β-amyloid levels (375, 376).