Ka Weng Leong
1,2
Mark M. W. Chong
1,2*- 1 RNA and T cell Biology, St Vincent’s Institute of Medical Research, Fitzroy, VIC, Australia
- 2 Department of Medicine, University of Melbourne, Parkville, VIC, Australia
The Microprocessor is an essential protein complex that is responsible for the first processing step in the biogenesis of canonical microRNAs. The core of this complex is composed of two proteins, the ribonuclease III enzyme DROSHA and its double-stranded RNA-binding cofactor DGCR8. Dysregulation of the expression of the Microprocessor contributes to many disorders, including pluripotency defects, immune dysfunction, cancers, and neurological diseases. Multiple post-translational modifications (PTMs) have been reported for DROSHA and DGCR8, and these are thought to play roles in regulating Microprocessor levels and its functions; however, most of these PTMs remain functionally uncharacterized. In this review, we discuss these PTMs of the Microprocessor, focusing on phosphorylation, acetylation, ubiquitination, and SUMOylation, and how these modifications are thought to regulate protein stability, microRNA production, and other non-canonical Microprocessor activities.
1 Introduction
MicroRNAs (miRNAs) are a class of small non-coding RNAs that play a critical role in gene regulation. They form complementary base-pairs with protein-coding messenger RNA (mRNA) targets, which lead to the degradation and translation repression of mRNAs (Bartel, 2018; Kim et al., 2025). Each miRNA can target multiple mRNAs (Lim et al., 2005), and collectively, miRNAs are predicted to regulate more than half of the protein-coding genes in the genome (Friedman et al., 2009). Tight regulation of miRNA biogenesis is crucial for maintaining the normal physiological state of cells, and dysregulation of miRNAs is associated with many diseases, such as developmental defects, autoimmune disorders, and cancer (Bartel, 2018; Chen et al., 2016; Garzon et al., 2006; Jeker et al., 2013; Volinia et al., 2006).
In canonical miRNA biogenesis, the primary (pri)-miRNAs are transcribed and co-transcriptionally processed by the Microprocessor complex (Morlando et al., 2008) (Figure 1A). It is minimally composed of the ribonuclease III (RNase III) protein DROSHA, and its double-stranded (ds)RNA-binding partner, DGCR8 (Nguyen et al., 2015). The complex recognizes and cleaves miRNA-containing stem-loop structures embedded within the long pri-miRNA transcripts (Morlando et al., 2008; Yin et al., 2015). These stem-loops, or pre-miRNAs, are then exported to the cytoplasm and further processed by another RNase III protein, DICER (Lee et al., 2002). The resulting miRNA duplex is loaded onto an argonaute (AGO) protein, the core component of the RNA-induced silencing complex (RISC), where one strand is retained as the mature miRNA (Kawamata and Tomari, 2010).
Figure 1. Post-translational modifications regulate the miRNA biogenesis machinery. Pri-miRNAs are transcribed by RNA polymerase II (Pol II) and co-transcriptionally processed by the Microprocessor complex, composed of DROSHA and DGCR8. The complex recognises and releases the stem-loop pre-miRNA intermediate, which is then exported to the cytoplasm. There, another RNase III protein, DICER, together with TRBP, removes the apical loops and loads one strand (the mature miRNA) into an AGO-containing RNA-induced silencing complex (RISC) to mediate mRNA degradation or translational repression. Components of the miRNA machinery, including DROSHA, DGCR8, TRBP, DICER, and AGO, are all regulated by post-translational modifications (PTMs). Summary of the enzymatic reactions that add/remove the four PTMs found on Microprocessor components. Kinases transfer a phosphate group (P) from ATP to serine, threonine, or tyrosine residues of target proteins, whereas phosphatases reverse the modification. Acetyltransferases transfer an acetyl group (Ac) from coenzyme A (CoA) to lysine residues of target proteins, while deacetylases reverse the modification. Ubiquitin-activating enzymes (E1), ubiquitin-conjugating enzymes (E2), and ubiquitin ligases (E3) sequentially facilitate the transfer of ubiquitin (Ub) to lysine residues of target proteins. Ubiquitin itself can be ubiquitinated, forming differently linked ubiquitin chains, such as K48-linked chains. Ubiquitination is reversed by deubiquitinases. SUMO-activating enzymes (E1), SUMO-conjugating enzymes (E2), and SUMO ligases (E3) successively catalyse the transfer of small ubiquitin-like modifier (SUMO) to lysine residues within the SUMOylation consensus motif ψKxD/E, where ψ is a large hydrophobic amino acid, x is any amino acid, D and E are acidic amino acids (aspartic acid and glutamic acid, respectively). Only SUMO1 (S1) and SUMO2 (S2) are shown in the illustration, while multiple SUMO isoforms exist. SUMO-specific proteases (SENPs) remove SUMO from SUMO-modified proteins. Created in BioRender (https://BioRender.com/98kt2ep).
In addition to microRNA biogenesis, the Microprocessor has also been reported to exert other molecular functions. These include recognizing stem-loop-containing viral RNAs (Aguado et al., 2017), binding or cleaving stem-loop-embedded protein-coding mRNAs (Cirera-Salinas et al., 2017; Han et al., 2009; Havens et al., 2014; Johanson et al., 2015; Knuckles et al., 2012; Lee et al., 2017; Marinaro et al., 2017; Rolando et al., 2016), and interacting with DNA repair proteins (Cabrini et al., 2021; Calses et al., 2017; Hang et al., 2021; Lu et al., 2018). Through these activities, the Microprocessor contributes to a range of other biological processes, including anti-viral defense (Aguado et al., 2017), autoregulation (Han et al., 2009; Lee et al., 2017), cell fate determination (Johanson et al., 2015; Knuckles et al., 2012; Marinaro et al., 2017; Rolando et al., 2016), alternative splicing (Cirera-Salinas et al., 2017; Havens et al., 2014; Lee et al., 2017), and DNA damage repair (Cabrini et al., 2021; Calses et al., 2017; Hang et al., 2021; Lu et al., 2018).
PTMs are the covalent addition or removal of modifying groups on the side chain of amino acid residues (Ramazi and Zahiri, 2021), and they can alter the chemical and physical properties of proteins, thereby regulating their subcellular localization, structures, stability, and functions (Ryšlavá et al., 2013). A growing number of PTMs have been reported for the miRNA machinery and their associated proteins, suggesting that PTMs are an important mechanism of regulating the miRNA pathway (Ha and Kim, 2014) (Figure 1A). This includes ERK phosphorylation and stabilization of TRBP, the cofactor of DICER, which promotes mature miRNA production (Paroo et al., 2009). Another is the SUMO2/3 modification of DICER, which downregulates protein levels and its processing of pre-miRNAs (Gross et al., 2014). Both Microprocessor proteins, DROSHA and DGCR8, also appear to be heavily modified by phosphorylation, acetylation, ubiquitination, and SUMOylation. DROSHA is also reportedly methylated at a single residue, R271, which was identified by high-throughput discovery mass spectrometry, but its functional relevance remains unknown (Larsen et al., 2016). In this review, we will summarize the reported phosphorylation, acetylation, ubiquitination, and SUMOylation of DROSHA and DGCR8 and discuss how these PTMs regulate the diverse functions of the Microprocessor (Table 1).
Table 1. PTMs identified on human DROSHA and DGCR8.
2 Implications of microprocessor dysregulation
DROSHA is a nuclear RNase III that contains two tandem RNase III domains (RIIIDs), RIIIDa and RIIIDb, followed by a dsRNA binding domain (dsRBD) at the C-terminus (Kwon et al., 2016). The N-terminal extension comprises a proline-rich (P-rich) domain and an arginine/serine-rich (R/S-rich) domain, followed by a central domain (CED) (Kwon et al., 2016). DGCR8 is a nuclear dsRNA-binding protein composed of an N-terminal tail, an RNA-binding heme domain (Rhed), followed by two dsRBDs and the C-terminal tail (CTT) (Senturia et al., 2010). The DROSHA dsRBDs alone have weak affinity for dsRNAs and require a dimer of DGCR8 for RNA recruitment and substrate recognition (Kranick et al., 2017; Roth et al., 2013; Wostenberg et al., 2010). DGCR8 stabilizes DROSHA by binding DROSHA RIIIDs with its CTT, while DROSHA controls DGCR8 expression by cleaving DGCR8 mRNA (Han et al., 2009; Nguyen et al., 2015).
Given the molecular functions of the Microprocessor, unsurprisingly, dysregulation of either DROSHA or DGCR8 can result in adverse consequences. Both proteins are critical for early embryogenesis. DROSHA or DGCR8 deficiency in mice results in impaired embryonic stem cell differentiation and lethality soon after implantation (Deng et al., 2019; Lyu et al., 2024; Wang et al., 2007). Loss of DROSHA (Chong et al., 2008) or DGCR8 (Jeker et al., 2013) disrupts miRNA production and impairs regulatory T cells homeostasis and thus is suggested to drive immune disorders. In humans, the loss of one functional allele of DGCR8 is sufficient to impair miRNA production, suggesting to be the cause of the physiological defects seen in 22q11.2 deletion syndrome, including pluripotency defects, immunodeficiency due to an underdeveloped thymus, and psychiatric illnesses (Colomer-Boronat et al., 2025; McDonald-McGinn et al., 2015).
Mutations in DGCR8 are observed in thyroid cancer (Paulsson et al., 2021; Rodrigues et al., 2024; Rodrigues et al., 2022), and mutations in both DROSHA and DGCR8 occur in Wilms’ tumor (Rakheja et al., 2014; Walz et al., 2015), highlighting the importance of maintaining Microprocessor integrity. While dysregulated expression of these Microprocessor components or the mislocalization of DROSHA have been reported in a range of other cancers (Huang et al., 2014; Lin and Gregory, 2015; Theotoki et al., 2025).
Neurological disorders can arise from abnormal intracellular localization or downregulation of Microprocessor components. In fragile X-associated tremor/ataxia syndrome, DROSHA and DGCR8 are sequestered by nuclear RNA aggregates (Sellier et al., 2013). While in amyotrophic lateral sclerosis, DROSHA is mislocalized into the cytoplasm and forms aggregates with the FUS mutant (Kour et al., 2023). In traumatic brain injury, phosphorylation of DROSHA is associated with downregulated expression, which drives neuronal apoptosis and further exacerbates the condition (Huang et al., 2024). DGCR8 deficiency is implicated in the impaired cognition and schizophrenia observed in 22q11.2 deletion syndrome (Fénelon et al., 2011; Ouchi et al., 2013; Rao et al., 2025).
These are examples of the consequences of the most severe lesions of DROSHA and DGCR8 deficiency. However, more nuanced phenotypes are likely to occur due to defective PTMs of these proteins.
3 Post-translational modifications on the microprocessor
3.1 Phosphorylation
Phosphorylation of a protein is the reversible transfer of a phosphoryl group (PO3 2-) from an adenosine triphosphate (ATP) to a specific residue by kinases, most commonly to the hydroxyl group-containing serine, threonine, and tyrosine (Ardito et al., 2017) (Figure 1B). This modification is reversed by phosphatases (Ardito et al., 2017). The addition of a phosphoryl group increases the polarity and hydrophilicity of a protein, affecting conformation and substrate affinity, and thus the functions of the protein (Ardito et al., 2017).
The R/S-rich domain in the N-terminal of DROSHA provides an extensive platform for phosphorylation events, with 14 out of 21 phosphosites documented in PhosphoSitePlus v6.8.1 located in this domain (Hornbeck et al., 2015). Glycogen synthase kinase 3 beta (GSK3β) and Polo-Like Kinase 1 (PLK1) have been reported to phosphorylate DROSHA at S300 and S302, which promotes its nuclear translocation, its association with DGCR8, and the downstream miRNA production (Fletcher et al., 2023; Tang et al., 2011; Tang et al., 2010). p38 mitogen-activated protein kinase (MAPK) reportedly phosphorylates another group of residues, S221, S255, T274, S300, and S355 in DROSHA, upon cellular stress (Yang et al., 2015). This drives its dissociation from DGCR8, translocation to the cytoplasm, and ultimately degradation by calpain (Yang et al., 2015). The consequence is a reduction in the miRNA biogenesis, leading to apoptosis (Yang et al., 2015). Interestingly, these examples demonstrate that the phosphorylation of S300 can facilitate counterposed translocations, though the nuclear export of DROSHA additionally requires the phosphorylation of several other residues in the domain. Besides p38 MAPK, CDK5 has also been reported to phosphorylate S221, S255, T274, S300, and S355 in DROSHA after traumatic brain injury, leading to its degradation by calpain and cell apoptosis (Huang et al., 2024).
A phosphoproteomic analysis of overexpressed human DGCR8 in insect or human cell lines identified 23 phosphorylated sites, with 5 of these sites phosphorylated only in the insect cells (Herbert et al., 2013). Mutating all 23 sites at once revealed that DGCR8 can be phosphorylated by extracellular signal-regulated kinase (ERK), which contributes to the stabilization of the protein, pro-growth miRNA production, and enhanced cell proliferation (Zhu et al., 2015). The phosphorylation of the 23 sites was later also reported to be important for controlling the alternative splicing of Tcf7l1 mRNA, and this serves as a checkpoint for stem cell differentiation (Cirera-Salinas et al., 2017). However, which specific phosphosites are important remained ambiguous. A separate study showed that epidermal growth factor-induced ERK activation also results in DGCR8 phosphorylation (Zhu et al., 2015). This phosphorylates S109, S153, T371, and S377, and was predicted to be responsible for promoting DGCR8 SUMOylation (Zhu et al., 2015). Concordantly, coilin, a nuclear Cajal bodies structural protein, was suggested to enhance S377 phosphorylation, which in turn promotes DGCR8 stability and the downstream miRNA biogenesis (Lett et al., 2021).
The phosphorylation of specific sites in DGCR8 were also reported to regulate DNA damage repair pathways in either a miRNA-dependent or a miRNA-independent manner. DNA repair mechanisms are crucial for preventing the accumulation of mutations and cellular transformation. The phosphorylation at S153 by Jun N-terminal kinase (JNK) confers resistance to mammalian cells against ultraviolet radiation through the transcription-coupled nucleotide excision repair (TC-NER) pathway (Calses et al., 2017). Upon ionizing radiation, DGCR8 S677 is phosphorylated by the kinase ATM, allowing it to be recognized by USP51, which enhances its protein stabilization (Hang et al., 2021). This phosphorylated DGCR8 can then contribute to double-stranded break repair by recruiting RNF168 to RNF8 and MDC1 for histone ubiquitination (Hang et al., 2021). The function of the phosphorylated DGCR8 in DNA repair after ultraviolet or ionizing radiation is independent of both DROSHA and miRNA (Calses et al., 2017; Hang et al., 2021). The tyrosine kinase ABL was also reported to phosphorylate DGCR8 at Y267, potentially in a tissue-specific manner, to reinforce the interaction between DGCR8 and its RNA substrate for the production of DROSHA-dependent miR-34c, which is involved in DNA repair after DNA damage induced by cisplatin treatment (Tu et al., 2015). These together suggest that proper phosphorylation of DGCR8 prevents mutation accumulation and thus suppresses cancer development.
3.2 Acetylation
Acetylation, the addition of an acetyl group (CH3CO−) to a residue, is mediated by acetyltransferases and removed by deacetylases (Narita et al., 2019) (Figure 1C). Although acetylation occurs more frequently on lysine residues, it can also occur on the N-terminal amino acid and on serine, threonine, and tyrosine residues (Xia et al., 2020). The addition of an acetyl group to lysine neutralizes the positive charge and thus can alter the structure, stability, and function of a protein (Narita et al., 2019). Moreover, acetylation often competes with ubiquitination for the same lysine residues (Narita et al., 2019).
DROSHA contains 13 lysine residues at the N-terminus that can undergo acetylation or ubiquitination to regulate protein stability (Tang et al., 2013). DROSHA was reported to be weakly acetylated by p300 and this can be completely removed by HDAC1, a histone deacetylase that co-immunoprecipitates with the Microprocessor in HEK293 cells and K562 cells (Wada et al., 2012). More specifically, DROSHA was reported to be acetylated at K382 and other lysine residues by several histone acetyltransferases, including p300, CBP, and GCN5, and this acetylation is enhanced upon inhibition of HDAC-mediated deacetylation, resulting in DROSHA protein stabilization (Tang et al., 2013).
DGCR8 is acetylated in its native form (Wada et al., 2012). It can be further acetylated by p300 and can be almost fully deacetylated by HDAC1 (Wada et al., 2012). All three conserved and indispensable lysine residues K561, K562, and K565 in the dsRBD of DGCR8 can be deacetylated by HDAC1, which enhances the affinity of DGCR8 for primary miRNA substrates and thus regulates the production of miRNAs (Acevedo et al., 2015; Kranick et al., 2017; Wada et al., 2012; Wostenberg et al., 2010). This suggests HDAC1 exerts dual effects on gene silencing, through conventional histone deacetylation and chromatin condensation to suppress transcription, as well as enhanced miRNA production via the non-canonical DGCR8 deacetylation for post-transcriptional repression (Dunaway and Pollock, 2022; Wada et al., 2012).
3.3 Ubiquitination
Ubiquitination is the reversible process of the addition of ubiquitin molecules to the side chain of an amino acid residue by a protein complex containing ubiquitin-activating enzyme (E1), ubiquitin-conjugating enzyme (E2), and ubiquitin ligase (E3) (Damgaard, 2021) (Figure 1D). Ubiquitin is a small peptide of 76 amino acids that acts as a tag on proteins for the regulation of several cellular processes, including cell signaling, protein trafficking, and DNA repair, either via proteolytic or non-proteolytic mechanisms (Damgaard, 2021). Like acetylation, ubiquitination can occur on multiple amino acids but more frequently occurs at lysine, which, as mentioned above, can compete with acetylation (Kelsall, 2022). Apart from the targeted proteins, ubiquitin itself can also be ubiquitinated on any of the seven internal lysine residues, K6, K11, K27, K29, K33, K48, and K63, forming different lysine-linked chains of ubiquitin (Damgaard, 2021). The linkage affects the conformation of the poly-ubiquitin chains and thus signals different downstream pathways (Damgaard, 2021). For example, K48-linked polyubiquitination canonically functions as a tag for protein degradation by the 26S proteasome, while K63-linked polyubiquitination typically contributes to endocytosis and cellular signaling (Sun and Zhang, 2022; Wang et al., 2012).
Human DROSHA contains 70 lysine residues, of which ubiquitination was detected at 21 by high-throughput proteomic discovery mass spectrometry (Hornbeck et al., 2015). DROSHA protein stability is regulated via the K48-linked ubiquitin-proteasome pathway, with one or more of the 13 lysine residues in the DROSHA N-terminus being the potential ubiquitination target (Tang et al., 2013). mTOR activation indirectly reduces DROSHA protein level through ubiquitin-mediated degradation and drives global miRNA reduction (Ye et al., 2015). The activation upregulates the E3 ubiquitin ligase MDM2 in p53-dependent and -independent manners, and MDM2 then ubiquitinates DROSHA for proteasome-mediated degradation (Ye et al., 2015). This results in the reduction of specific miRNAs, including miR-297 and miR-567, to drive apoptosis (Ye et al., 2015). Whereas the deprivation of either amino acids or glucose reduces mTOR activation, resulting in reduced MDM2 level, stabilization of DROSHA, and thus improved survival (Ye et al., 2015). These suggest DROSHA plays a protective role under nutrient- and energy-restricted conditions and restricts cell overgrowth when there is sufficient resource.
Similarly, DGCR8 protein stability is regulated by the K48-linked ubiquitin-proteasomal pathway. DGCR8 contains 61 lysine residues. Although the basal level of DGCR8 ubiquitination was barely detectable (Li et al., 2023; Zhu et al., 2015), 13 of the lysine residues were reported to be ubiquitinated (Hornbeck et al., 2015). The proteasome inhibitor MG132, but not the lysosome inhibitor chloroquine, could block the degradation of DGCR8 (Zhu et al., 2015). Concordantly, as previously mentioned, DGCR8 is phosphorylated at S677 upon ionizing radiation for its recognition by USP51, a deubiquitinase, to remove the K48-linked ubiquitin for its stabilization (Hang et al., 2021).
3.4 SUMOylation
SUMOylation is another reversible process in which a small ubiquitin-like modifier (SUMO) molecule is covalently added to proteins, usually at a lysine residue embedded within a consensus motif (Celen and Sahin, 2020) (Figure 1E). The conjugatable SUMO family in mammals includes SUMO1, SUMO2, and SUMO3, which are around 100 amino acids and are structurally similar to ubiquitin (Han et al., 2018; Seeler and Dejean, 2003; Wang and Matunis, 2023). SUMO1 shares approximately 50% sequence identity with SUMO2 and SUMO3, while SUMO2 and SUMO3 are around 97% identical, making them difficult to discriminate from each other via antibody detection and thus are often referred to as SUMO2/3 (Pichler et al., 2017). SUMO proteins serve as regulatory tags that modulate target proteins for protein localization, stability, and molecular interactions, with SUMO1 primarily functioning under normal physiological conditions and SUMO2/3 under stressed conditions (Han et al., 2018; Seeler and Dejean, 2003). Like ubiquitination, SUMOylation is catalyzed by a cascade of enzymes, including a SUMO-activating enzyme (E1), a SUMO-conjugating enzyme (E2), and a SUMO ligase (E3), with the removal catalyzed by SUMO-specific proteases (SENPs) (Han et al., 2018). SUMOylation can be induced by phosphorylation and in turn can also affect other subsequent PTMs, including acetylation and ubiquitination (Wei et al., 2024). Similar to acetylation, SUMOylation can compete with ubiquitination for the same lysine residues to regulate protein stability (Wei et al., 2024).
DROSHA has been reported to be SUMOylated at K388 and K409 in a HeLa cell high-throughput dataset (Hendriks et al., 2014); however, the function of the SUMOylations remains unknown. DGCR8, on the other hand, has been reported to be subjected to SUMOylation at K259, K426, and K707, with K259 and K707 being the primary sites of SUMO attachment (Li et al., 2023; Zhu et al., 2016; 2015). SUMO1-conjugation at DGCR8 K259 prevents DGCR8 from nuclear export and confers tumor suppression function (Zhu et al., 2016). This modification is promoted by the tumor suppressor protein p14ARF that primarily functions by binding and sequestering MDM2 to prevent the downstream MDM2-mediated protein degradation (Cilluffo et al., 2020; Weber et al., 1999). Conversely, SUMO1 modification at DGCR8 K707 promotes tumorigenesis and metastasis (Zhu et al., 2015). This SUMOylation is promoted by the ERK-mediated DGCR8 phosphorylation, antagonizing DGCR8 ubiquitination and thus enhancing its protein stability (Zhu et al., 2015). It also reinforces the affinity of DGCR8 for its pri-miRNA substrates without affecting downstream processing (Zhu et al., 2015). DGCR8 can also be modified by SUMO2 at all three SUMO-susceptible lysine residues, with the SUMOylation mediated by the deubiquitinase USP36 (Li et al., 2023). USP36-mediated SUMOylation promotes DGCR8’s affinity towards its pri-miRNA substrates, as well as promotes cell proliferation without altering its expression and its interaction with DROSHA (Li et al., 2023).
4 Conclusions and perspectives
4.1 Summary of key insights
MiRNAs are important regulators of gene expression, facilitating different cellular processes through mediating mRNA degradation or translational repression. Consistent with their importance, their synthesis is tightly regulated at multiple levels, including transcription, splicing, and RNA modifications (Kim et al., 2025). Beyond the transcriptional regulation of the miRNA genes themselves, the miRNA machinery and their associated proteins are also strictly regulated to fine-tune the miRNA production, with PTMs playing an essential role in the modulation.
In this review, we have summarized the reported PTMs of the Microprocessor, which modulate the Microprocessor in multiple ways, from its expression to substrate affinity, as well as its protein-protein interactions. These PTMs can profoundly influence the canonical miRNA biogenesis and other non-canonical Microprocessor activities, including mRNA cleavage, mRNA alternative splicing, and DNA repair. Disruption of DROSHA or DGCR8 expression impairs miRNA production and contributes to developmental defects, immune disorders, and neurological diseases, while upregulation of DROSHA and DGCR8 can enhance proliferation, promoting tumorigenesis.
4.2 Current limitations and challenges
Although significant progress has been made in identifying PTMs, most reported sites have been identified primarily through high-throughput proteomic discovery mass spectrometry, with little or no functional investigation. Moreover, many studies often rely on the overexpression of tagged proteins in cell lines, and thus, identified PTMs may not necessarily be physiologically relevant. Where functions of specific PTMs have been assessed, these again have often been studied in cell lines. In the future, it will be important to address function by genetic manipulation of the endogenous proteins and to study the function of these PTMs in vivo, such as with mouse models.
Another challenge is that most PTMs are reversible and highly dynamic, making them difficult to be detected in real time. For instance, K48-linked ubiquitinated proteins are rapidly degraded via the proteasomal pathway and thus are hard to be captured and have the corresponding modified sites identified. Moreover, the SUMO1 DGCR8 SUMOylation examples indicate that the same type of modification can exert counteracting effects. Whereas the phosphorylation of DROSHA S300 participates in different phosphorylation patterns to control opposing translocations, highlighting the site-specific and collaborative nature of PTM regulation. Furthermore, PTMs sometimes function sequentially or even antagonistically. For example, DGCR8 phosphorylation facilitates SUMOylation; acetylation and ubiquitination compete on the same lysine residues in DROSHA. These complications make functional characterization technically challenging with current PTM-mimetic and -deficient mutant approaches, which can barely resolve the complicated crosstalk between PTMs.
4.3 Future directions
Addressing these challenges will require more sophisticated approaches and efforts to identify Microprocessor PTMs, elucidate their functions, and dissect the complexitiy of crosstalk between specific PTMs. Targeting PTMs on the Microprocessor or the upstream enzymes may provide strategies to restore the abnormal expression level and activity of the Microprocessor in diseases and potentially alleviate disease progression. Together, these efforts will deepen our understanding of RNA biology and offer novel therapeutic opportunities for disease diagnosis and intervention.
Author contributions
KL: Conceptualization, Writing – original draft, Writing – review and editing. MC: Conceptualization, Funding acquisition, Writing – review and editing.
Funding
The authors declare that financial support was received for the research and/or publication of this article. Research in the Chong laboratory is supported by grants from the National Health Medical Research Council (1117154 & 1122395), the US Department of Defense Lupus (W81XWH-19-1-0728), Breakthrough T1D (3-SRA-2024-1463-S-B), Diabetes Australia (Y20G-CHOM), mRNA Victoria and Cancer Research Institute.
Conflict of interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
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References
Acevedo, R., Orench-Rivera, N., Quarles, K. A., and Showalter, S. A. (2015). Helical defects in MicroRNA influence protein binding by TAR RNA binding protein. PLoS ONE 10, e0116749. doi:10.1371/journal.pone.0116749
Aguado, L. C., Schmid, S., May, J., Sabin, L. R., Panis, M., Blanco-Melo, D., et al. (2017). RNase III nucleases from diverse kingdoms serve as antiviral effectors. Nature 547, 114–117. doi:10.1038/nature22990
Akimov, V., Barrio-Hernandez, I., Hansen, S. V. F., Hallenborg, P., Pedersen, A.-K., Bekker-Jensen, D. B., et al. (2018). UbiSite approach for comprehensive mapping of lysine and N-terminal ubiquitination sites. Nat. Struct. Mol. Biol. 25, 631–640. doi:10.1038/s41594-018-0084-y
Ardito, F., Giuliani, M., Perrone, D., Troiano, G., and Lo Muzio, L. (2017). The crucial role of protein phosphorylation in cell signaling and its use as targeted therapy (review). Int. J. Mol. Med. 40, 271–280. doi:10.3892/ijmm.2017.3036
Beli, P., Lukashchuk, N., Wagner, S. A., Weinert, B. T., Olsen, J. V., Baskcomb, L., et al. (2012). Proteomic investigations reveal a role for RNA processing factor THRAP3 in the DNA damage response. Mol. Cell 46, 212–225. doi:10.1016/j.molcel.2012.01.026
Boeing, S., Williamson, L., Encheva, V., Gori, I., Saunders, R. E., Instrell, R., et al. (2016). Multiomic analysis of the UV-induced DNA damage response. Cell Rep. 15, 1597–1610. doi:10.1016/j.celrep.2016.04.047
Cabrini, M., Roncador, M., Galbiati, A., Cipolla, L., Maffia, A., Iannelli, F., et al. (2021). DROSHA is recruited to DNA damage sites by the MRN complex to promote non-homologous end joining. J. Cell Sci. 134, jcs249706. doi:10.1242/jcs.249706
Calses, P. C., Dhillon, K. K., Tucker, N., Chi, Y., Huang, J.-W., Kawasumi, M., et al. (2017). DGCR8 mediates repair of UV-induced DNA damage independently of RNA processing. Cell Rep. 19, 162–174. doi:10.1016/j.celrep.2017.03.021
Celen, A. B., and Sahin, U. (2020). Sumoylation on its 25th anniversary: mechanisms, pathology, and emerging concepts. FEBS J. 287, 3110–3140. doi:10.1111/febs.15319
Chen, J.-Q., Papp, G., Szodoray, P., and Zeher, M. (2016). The role of microRNAs in the pathogenesis of autoimmune diseases. Autoimmun. Rev. 15, 1171–1180. doi:10.1016/j.autrev.2016.09.003
Chong, M. M., Rasmussen, J. P., Rudensky, A. Y., and Littman, D. R. (2008). The RNAseIII enzyme DROSHA is critical in T cells for preventing lethal inflammatory disease. J. Exp. Med. 205, 2005–2017. doi:10.1084/jem.20081219
Christensen, G. L., Kelstrup, C. D., Lyngsø, C., Sarwar, U., Bøgebo, R., Sheikh, S. P., et al. (2010). Quantitative phosphoproteomics dissection of seven-transmembrane receptor signaling using full and biased agonists. Mol. Cell. Proteomics MCP 9, 1540–1553. doi:10.1074/mcp.M900550-MCP200
Cilluffo, D., Barra, V., and Di Leonardo, A. (2020). P14ARF: the absence that makes the difference. Genes 11, 824. doi:10.3390/genes11070824
Cirera-Salinas, D., Yu, J., Bodak, M., Ngondo, R. P., Herbert, K. M., and Ciaudo, C. (2017). Noncanonical function of DGCR8 controls mESC exit from pluripotency. J. Cell Biol. 216, 355–366. doi:10.1083/jcb.201606073
Colomer-Boronat, A., Knol, L. I., Peris, G., Sanchez, L., Peluso, S., Tristan-Ramos, P., et al. (2025). DGCR8 haploinsufficiency leads to primate-specific RNA dysregulation and pluripotency defects. Nucleic Acids Res. 53, gkaf197. doi:10.1093/nar/gkaf197
Damgaard, R. B. (2021). The ubiquitin system: from cell signalling to disease biology and new therapeutic opportunities. Cell Death Differ. 28, 423–426. doi:10.1038/s41418-020-00703-w
Deng, L., Ren, R., Liu, Z., Song, M., Li, J., Wu, Z., et al. (2019). Stabilizing heterochromatin by DGCR8 alleviates senescence and osteoarthritis. Nat. Commun. 10, 3329. doi:10.1038/s41467-019-10831-8
Dunaway, L. S., and Pollock, J. S. (2022). HDAC1: an environmental sensor regulating endothelial function. Cardiovasc. Res. 118, 1885–1903. doi:10.1093/cvr/cvab198
Fénelon, K., Mukai, J., Xu, B., Hsu, P.-K., Drew, L. J., Karayiorgou, M., et al. (2011). Deficiency of Dgcr8, a gene disrupted by the 22q11.2 microdeletion, results in altered short-term plasticity in the prefrontal cortex. Proc. Natl. Acad. Sci. U. S. A. 108, 4447–4452. doi:10.1073/pnas.1101219108
Fletcher, C. E., Taylor, M. A., and Bevan, C. L. (2023). PLK1 regulates MicroRNA biogenesis through DROSHA phosphorylation. Int. J. Mol. Sci. 24, 14290. doi:10.3390/ijms241814290
Franz-Wachtel, M., Eisler, S. A., Krug, K., Wahl, S., Carpy, A., Nordheim, A., et al. (2012). Global detection of protein kinase D-dependent phosphorylation events in nocodazole-treated human cells. Mol. Cell. Proteomics MCP 11, 160–170. doi:10.1074/mcp.M111.016014
Friedman, R. C., Farh, K.K.-H., Burge, C. B., and Bartel, D. P. (2009). Most mammalian mRNAs are conserved targets of microRNAs. Genome Res. 19, 92–105. doi:10.1101/gr.082701.108
Garzon, R., Fabbri, M., Cimmino, A., Calin, G. A., and Croce, C. M. (2006). MicroRNA expression and function in cancer. Trends Mol. Med. 12, 580–587. doi:10.1016/j.molmed.2006.10.006
Gross, T. J., Powers, L. S., Boudreau, R. L., Brink, B., Reisetter, A., Goel, K., et al. (2014). A microRNA processing defect in smokers’ macrophages is linked to SUMOylation of the endonuclease DICER. J. Biol. Chem. 289, 12823–12834. doi:10.1074/jbc.M114.565473
Grosstessner-Hain, K., Hegemann, B., Novatchkova, M., Rameseder, J., Joughin, B. A., Hudecz, O., et al. (2011). Quantitative phospho-proteomics to investigate the polo-like kinase 1-dependent phospho-proteome. Mol. Cell. Proteomics 10, M111.008540. doi:10.1074/mcp.M111.008540
Ha, M., and Kim, V. N. (2014). Regulation of microRNA biogenesis. Nat. Rev. Mol. Cell Biol. 15, 509–524. doi:10.1038/nrm3838
Han, J., Pedersen, J. S., Kwon, S. C., Belair, C. D., Kim, Y.-K., Yeom, K.-H., et al. (2009). Posttranscriptional crossregulation between DROSHA and DGCR8. Cell 136, 75–84. doi:10.1016/j.cell.2008.10.053
Han, Z.-J., Feng, Y.-H., Gu, B.-H., Li, Y.-M., and Chen, H. (2018). The post-translational modification, SUMOylation, and cancer (review). Int. J. Oncol. 52, 1081–1094. doi:10.3892/ijo.2018.4280
Hang, Q., Zeng, L., Wang, L., Nie, L., Yao, F., Teng, H., et al. (2021). Non-canonical function of DGCR8 in DNA double-strand break repair signaling and tumor radioresistance. Nat. Commun. 12, 4033. doi:10.1038/s41467-021-24298-z
Havens, M. A., Reich, A. A., and Hastings, M. L. (2014). DROSHA promotes splicing of a pre-microRNA-like alternative exon. PLoS Genet. 10, e1004312. doi:10.1371/journal.pgen.1004312
Hendriks, I. A., D’Souza, R. C. J., Yang, B., Verlaan-de Vries, M., Mann, M., and Vertegaal, A. C. O. (2014). Uncovering global SUMOylation signaling networks in a site-specific manner. Nat. Struct. Mol. Biol. 21, 927–936. doi:10.1038/nsmb.2890
Herbert, K. M., Pimienta, G., DeGregorio, S. J., Alexandrov, A., and Steitz, J. A. (2013). Phosphorylation of DGCR8 increases its intracellular stability and induces a progrowth miRNA profile. Cell Rep. 5, 1070–1081. doi:10.1016/j.celrep.2013.10.017
Hornbeck, P. V., Zhang, B., Murray, B., Kornhauser, J. M., Latham, V., and Skrzypek, E. (2015). PhosphoSitePlus, 2014: mutations, PTMs and recalibrations. Nucleic Acids Res. 43, D512–D520. doi:10.1093/nar/gku1267
Huang, J.-T., Wang, J., Srivastava, V., Sen, S., and Liu, S.-M. (2014). MicroRNA machinery genes as novel biomarkers for cancer. Front. Oncol. 4, 113. doi:10.3389/fonc.2014.00113
Huang, L., Xia, L., Nie, T., Cui, B., Lu, J., Lu, F., et al. (2024). Maintaining DROSHA expression with Cdk5 inhibitors as a potential therapeutic strategy for early intervention after TBI. Exp. Mol. Med. 56, 210–219. doi:10.1038/s12276-023-01152-4
Jeker, L. T., Zhou, X., Blelloch, R., and Bluestone, J. A. (2013). DGCR8-mediated production of canonical micrornas is critical for regulatory T cell function and stability. PLoS ONE 8, e66282. doi:10.1371/journal.pone.0066282
Johanson, T. M., Keown, A. A., Cmero, M., Yeo, J. H. C., Kumar, A., Lew, A. M., et al. (2015). DROSHA controls dendritic cell development by cleaving messenger RNAs encoding inhibitors of myelopoiesis. Nat. Immunol. 16, 1134–1141. doi:10.1038/ni.3293
Kawamata, T., and Tomari, Y. (2010). Making RISC. Trends Biochem. Sci. 35, 368–376. doi:10.1016/j.tibs.2010.03.009
Kelsall, I. R. (2022). Non-lysine ubiquitylation: doing things differently. Front. Mol. Biosci. 9, 1008175. doi:10.3389/fmolb.2022.1008175
Kettenbach, A. N., Schweppe, D. K., Faherty, B. K., Pechenick, D., Pletnev, A. A., and Gerber, S. A. (2011). Quantitative phosphoproteomics identifies substrates and functional modules of Aurora and Polo-like kinase activities in mitotic cells. Sci. Signal. 4, rs5. doi:10.1126/scisignal.2001497
Kim, H., Lee, Y.-Y., and Kim, V. N. (2025). The biogenesis and regulation of animal microRNAs. Nat. Rev. Mol. Cell Biol. 26, 276–296. doi:10.1038/s41580-024-00805-0
Klammer, M., Kaminski, M., Zedler, A., Oppermann, F., Blencke, S., Marx, S., et al. (2012). Phosphosignature predicts dasatinib response in non-small cell lung cancer. Mol. Cell. Proteomics MCP 11, 651–668. doi:10.1074/mcp.M111.016410
Knuckles, P., Vogt, M. A., Lugert, S., Milo, M., Chong, M. M. W., Hautbergue, G. M., et al. (2012). DROSHA regulates neurogenesis by controlling neurogenin 2 expression independent of microRNAs. Nat. Neurosci. 15, 962–969. doi:10.1038/nn.3139
Kour, S., Fortuna, T., Anderson, E. N., Mawrie, D., Bilstein, J., Sivasubramanian, R., et al. (2023). DROSHA-dependent microRNAs modulate FUS-mediated neurodegeneration in vivo. Nucleic Acids Res. 51, 11258–11276. doi:10.1093/nar/gkad774
Kranick, J. C., Chadalavada, D. M., Sahu, D., and Showalter, S. A. (2017). Engineering double-stranded RNA binding activity into the DROSHA double-stranded RNA binding domain results in a loss of microRNA processing function. PLoS ONE 12, e0182445. doi:10.1371/journal.pone.0182445
Kwon, S. C., Nguyen, T. A., Choi, Y.-G., Jo, M. H., Hohng, S., Kim, V. N., et al. (2016). Structure of human DROSHA. Cell 164, 81–90. doi:10.1016/j.cell.2015.12.019
Larsen, S. C., Sylvestersen, K. B., Mund, A., Lyon, D., Mullari, M., Madsen, M. V., et al. (2016). Proteome-wide analysis of arginine monomethylation reveals widespread occurrence in human cells. Sci. Signal. 9, rs9. doi:10.1126/scisignal.aaf7329
Lee, Y., Jeon, K., Lee, J.-T., Kim, S., and Kim, V. N. (2002). MicroRNA maturation: stepwise processing and subcellular localization. EMBO J. 21, 4663–4670. doi:10.1093/emboj/cdf476
Lee, D., Nam, J.-W., and Shin, C. (2017). DROSHA targets its own transcript to modulate alternative splicing. RNA N. Y. N. 23, 1035–1047. doi:10.1261/rna.059808.116
Lett, K. E., Logan, M. K., McLaurin, D. M., and Hebert, M. D. (2021). Coilin enhances phosphorylation and stability of DGCR8 and promotes miRNA biogenesis. Mol. Biol. Cell 32, br4. doi:10.1091/mbc.E21-05-0225
Li, Y., Carey, T. S., Feng, C. H., Zhu, H.-M., Sun, X.-X., and Dai, M.-S. (2023). The ubiquitin-specific protease USP36 associates with the microprocessor complex and regulates miRNA biogenesis by SUMOylating DGCR8. Cancer Res. Commun. 3, 459–470. doi:10.1158/2767-9764.CRC-22-0344
Lim, L. P., Lau, N. C., Garrett-Engele, P., Grimson, A., Schelter, J. M., Castle, J., et al. (2005). Microarray analysis shows that some microRNAs downregulate large numbers of target mRNAs. Nature 433, 769–773. doi:10.1038/nature03315
Lin, S., and Gregory, R. I. (2015). MicroRNA biogenesis pathways in cancer. Nat. Rev. Cancer 15, 321–333. doi:10.1038/nrc3932
Lu, W.-T., Hawley, B. R., Skalka, G. L., Baldock, R. A., Smith, E. M., Bader, A. S., et al. (2018). DROSHA drives the formation of DNA:RNA hybrids around DNA break sites to facilitate DNA repair. Nat. Commun. 9, 532. doi:10.1038/s41467-018-02893-x
Lyu, X., Cui, Y., Kong, Y., Yang, M., Shen, H., Liao, S., et al. (2024). A transient transcriptional activation governs unpolarized-to-polarized morphogenesis during embryo implantation. Mol. Cell 84, 2665–2681.e13. doi:10.1016/j.molcel.2024.06.005
Marinaro, F., Marzi, M. J., Hoffmann, N., Amin, H., Pelizzoli, R., Niola, F., et al. (2017). MicroRNA-independent functions of DGCR8 are essential for neocortical development and TBR1 expression. EMBO Rep. 18, 603–618. doi:10.15252/embr.201642800
McDonald-McGinn, D. M., Sullivan, K. E., Marino, B., Philip, N., Swillen, A., Vorstman, J. A. S., et al. (2015). 22q11.2 deletion syndrome. Nat. Rev. Dis. Primer 1, 15071. doi:10.1038/nrdp.2015.71
Mertins, P., Qiao, J. W., Patel, J., Udeshi, N. D., Clauser, K. R., Mani, D. R., et al. (2013). Integrated proteomic analysis of post-translational modifications by serial enrichment. Nat. Methods 10, 634–637. doi:10.1038/nmeth.2518
Mertins, P., Yang, F., Liu, T., Mani, D. R., Petyuk, V. A., Gillette, M. A., et al. (2014). Ischemia in tumors induces early and sustained phosphorylation changes in stress kinase pathways but does not affect global protein levels. Mol. Cell. Proteomics MCP 13, 1690–1704. doi:10.1074/mcp.M113.036392
Mertins, P., Mani, D. R., Ruggles, K. V., Gillette, M. A., Clauser, K. R., Wang, P., et al. (2016). Proteogenomics connects somatic mutations to signalling in breast cancer. Nature 534, 55–62. doi:10.1038/nature18003
Morlando, M., Ballarino, M., Gromak, N., Pagano, F., Bozzoni, I., and Proudfoot, N. J. (2008). Primary microRNA transcripts are processed co-transcriptionally. Nat. Struct. Mol. Biol. 15, 902–909. doi:10.1038/nsmb.1475
Narita, T., Weinert, B. T., and Choudhary, C. (2019). Functions and mechanisms of non-histone protein acetylation. Nat. Rev. Mol. Cell Biol. 20, 156–174. doi:10.1038/s41580-018-0081-3
Nguyen, T. A., Jo, M. H., Choi, Y.-G., Park, J., Kwon, S. C., Hohng, S., et al. (2015). Functional anatomy of the human microprocessor. Cell 161, 1374–1387. doi:10.1016/j.cell.2015.05.010
Olsen, J. V., Vermeulen, M., Santamaria, A., Kumar, C., Miller, M. L., Jensen, L. J., et al. (2010). Quantitative phosphoproteomics reveals widespread full phosphorylation site occupancy during mitosis. Sci. Signal. 3, ra3. doi:10.1126/scisignal.2000475
Ouchi, Y., Banno, Y., Shimizu, Y., Ando, S., Hasegawa, H., Adachi, K., et al. (2013). Reduced adult hippocampal neurogenesis and working memory deficits in the Dgcr8-deficient mouse model of 22q11.2 deletion-associated schizophrenia can be rescued by IGF2. J. Neurosci. 33, 9408–9419. doi:10.1523/JNEUROSCI.2700-12.2013
Pan, C., Olsen, J. V., Daub, H., and Mann, M. (2009). Global effects of kinase inhibitors on signaling networks revealed by quantitative phosphoproteomics. Mol. Cell. Proteomics MCP 8, 2796–2808. doi:10.1074/mcp.M900285-MCP200
Paroo, Z., Ye, X., Chen, S., and Liu, Q. (2009). Phosphorylation of the human microRNA-generating complex mediates MAPK/Erk signaling. Cell 139, 112–122. doi:10.1016/j.cell.2009.06.044
Paulsson, J. O., Rafati, N., DiLorenzo, S., Chen, Y., Haglund, F., Zedenius, J., et al. (2021). Whole-genome sequencing of follicular thyroid carcinomas reveal recurrent mutations in MicroRNA processing subunit DGCR8. J. Clin. Endocrinol. Metab. 106, 3265–3282. doi:10.1210/clinem/dgab471
Pichler, A., Fatouros, C., Lee, H., and Eisenhardt, N. (2017). SUMO conjugation - a mechanistic view. Biomol. Concepts 8, 13–36. doi:10.1515/bmc-2016-0030
Rakheja, D., Chen, K. S., Liu, Y., Shukla, A. A., Schmid, V., Chang, T.-C., et al. (2014). Somatic mutations in DROSHA and DICER1 impair microRNA biogenesis through distinct mechanisms in Wilms tumours. Nat. Commun. 2, 4802. doi:10.1038/ncomms5802
Ramazi, S., and Zahiri, J. (2021). Post-translational modifications in proteins: resources, tools and prediction methods. Database J. Biol. Databases Curation 2021, baab012. doi:10.1093/database/baab012
Rao, S. B., Sun, Z., Brundu, F., Chen, Y., Sun, Y., Zhu, H., et al. (2025). Aberrant pace of cortical neuron development in brain organoids from patients with 22q11.2 deletion syndrome-associated schizophrenia. Nat. Commun. 16, 6986. doi:10.1038/s41467-025-62187-x
Rigbolt, K. T. G., Prokhorova, T. A., Akimov, V., Henningsen, J., Johansen, P. T., Kratchmarova, I., et al. (2011). System-wide temporal characterization of the proteome and phosphoproteome of human embryonic stem cell differentiation. Sci. Signal. 4, rs3. doi:10.1126/scisignal.2001570
Rodrigues, L., Canberk, S., Macedo, S., Soares, P., and Vinagre, J. (2022). DGCR8 microprocessor subunit mutation and expression deregulation in thyroid lesions. Int. J. Mol. Sci. 23, 14812. doi:10.3390/ijms232314812
Rodrigues, L., Da Cruz Paula, A., Soares, P., and Vinagre, J. (2024). Unraveling the significance of DGCR8 and miRNAs in thyroid carcinoma. Cells 13, 561. doi:10.3390/cells13070561
Rolando, C., Erni, A., Grison, A., Beattie, R., Engler, A., Gokhale, P. J., et al. (2016). Multipotency of adult hippocampal NSCs in vivo is restricted by DROSHA/NFIB. Cell Stem Cell 19, 653–662. doi:10.1016/j.stem.2016.07.003
Roth, B. M., Ishimaru, D., and Hennig, M. (2013). The core microprocessor component DiGeorge syndrome critical region 8 (DGCR8) is a nonspecific RNA-binding protein. J. Biol. Chem. 288, 26785–26799. doi:10.1074/jbc.M112.446880
Ryšlavá, H., Doubnerová, V., Kavan, D., and Vaněk, O. (2013). Effect of posttranslational modifications on enzyme function and assembly. J. Proteomics 92, 80–109. doi:10.1016/j.jprot.2013.03.025
Santamaria, A., Wang, B., Elowe, S., Malik, R., Zhang, F., Bauer, M., et al. (2011). The Plk1-dependent phosphoproteome of the early mitotic spindle. Mol. Cell. Proteomics MCP 10, M110.004457. doi:10.1074/mcp.M110.004457
Schreiber, T. B., Mäusbacher, N., Kéri, G., Cox, J., and Daub, H. (2010). An integrated phosphoproteomics work flow reveals extensive network regulation in early lysophosphatidic acid signaling. Mol. Cell. Proteomics MCP 9, 1047–1062. doi:10.1074/mcp.M900486-MCP200
Schweppe, D. K., Rigas, J. R., and Gerber, S. A. (2013). Quantitative phosphoproteomic profiling of human non-small cell lung cancer tumors. J. Proteomics 91, 286–296. doi:10.1016/j.jprot.2013.07.023
Seeler, J.-S., and Dejean, A. (2003). Nuclear and unclear functions of SUMO. Nat. Rev. Mol. Cell Biol. 4, 690–699. doi:10.1038/nrm1200
Sellier, C., Freyermuth, F., Tabet, R., Tran, T., He, F., Ruffenach, F., et al. (2013). Sequestration of DROSHA and DGCR8 by expanded CGG RNA repeats alters MicroRNA processing in fragile X-associated tremor/ataxia syndrome. Cell Rep. 3, 869–880. doi:10.1016/j.celrep.2013.02.004
Senturia, R., Faller, M., Yin, S., Loo, J. A., Cascio, D., Sawaya, M. R., et al. (2010). Structure of the dimerization domain of DiGeorge critical region 8: the dimerization domain of DGCR8. Protein Sci. 19, 1354–1365. doi:10.1002/pro.414
Sharma, K., D’Souza, R. C. J., Tyanova, S., Schaab, C., Wiśniewski, J. R., Cox, J., et al. (2014). Ultradeep human phosphoproteome reveals a distinct regulatory nature of Tyr and Ser/Thr-based signaling. Cell Rep. 8, 1583–1594. doi:10.1016/j.celrep.2014.07.036
Shiromizu, T., Adachi, J., Watanabe, S., Murakami, T., Kuga, T., Muraoka, S., et al. (2013). Identification of missing proteins in the neXtProt database and unregistered phosphopeptides in the PhosphoSitePlus database as part of the chromosome-centric human proteome project. J. Proteome Res. 12, 2414–2421. doi:10.1021/pr300825v
Stuart, S. A., Houel, S., Lee, T., Wang, N., Old, W. M., and Ahn, N. G. (2015). A phosphoproteomic comparison of B-RAFV600E and MKK1/2 inhibitors in melanoma cells. Mol. Cell. Proteomics MCP 14, 1599–1615. doi:10.1074/mcp.M114.047233
Sui, S., Wang, J., Yang, B., Song, L., Zhang, J., Chen, M., et al. (2008). Phosphoproteome analysis of the human Chang liver cells using SCX and a complementary mass spectrometric strategy. Proteomics 8, 2024–2034. doi:10.1002/pmic.200700896
Sun, M., and Zhang, X. (2022). Current methodologies in protein ubiquitination characterization: from ubiquitinated protein to ubiquitin chain architecture. Cell Biosci. 12, 126. doi:10.1186/s13578-022-00870-y
Tang, X., Zhang, Y., Tucker, L., and Ramratnam, B. (2010). Phosphorylation of the RNase III enzyme DROSHA at Serine300 or Serine302 is required for its nuclear localization. Nucleic Acids Res. 38, 6610–6619. doi:10.1093/nar/gkq547
Tang, X., Li, M., Tucker, L., and Ramratnam, B. (2011). Glycogen synthase kinase 3 beta (GSK3β) phosphorylates the RNAase III enzyme DROSHA at S300 and S302. PLoS One 6, e20391. doi:10.1371/journal.pone.0020391
Tang, X., Wen, S., Zheng, D., Tucker, L., Cao, L., Pantazatos, D., et al. (2013). Acetylation of DROSHA on the N-terminus inhibits its degradation by ubiquitination. PLoS ONE 8, e72503. doi:10.1371/journal.pone.0072503
Theotoki, E. I., Kakoulidis, P., Papavassiliou, K. A., Nikolakopoulos, K.-S., Vlachou, E. N., Basdra, E. K., et al. (2025). Non-canonical compartmentalization of DROSHA protein at the golgi apparatus: miRNA biogenesis-independent functionality in human cancer cells of diverse tissue origin. Int. J. Mol. Sci. 26, 9319. doi:10.3390/ijms26199319
Tu, C.-C., Zhong, Y., Nguyen, L., Tsai, A., Sridevi, P., Tarn, W. Y., et al. (2015). The kinase ABL phosphorylates the microprocessor subunit DGCR8 to stimulate primary microRNA processing in response to DNA damage. Sci. Signal. 8, ra64. doi:10.1126/scisignal.aaa4468
Udeshi, N. D., Svinkina, T., Mertins, P., Kuhn, E., Mani, D. R., Qiao, J. W., et al. (2013). Refined preparation and use of anti-diglycine remnant (K-ε-GG) antibody enables routine quantification of 10,000s of ubiquitination sites in single proteomics experiments. Mol. Cell. Proteomics MCP 12, 825–831. doi:10.1074/mcp.O112.027094
Van Hoof, D., Muñoz, J., Braam, S. R., Pinkse, M. W. H., Linding, R., Heck, A. J. R., et al. (2009). Phosphorylation dynamics during early differentiation of human embryonic stem cells. Cell Stem Cell 5, 214–226. doi:10.1016/j.stem.2009.05.021
Volinia, S., Calin, G. A., Liu, C.-G., Ambs, S., Cimmino, A., Petrocca, F., et al. (2006). A microRNA expression signature of human solid tumors defines cancer gene targets. Proc. Natl. Acad. Sci. U. S. A. 103, 2257–2261. doi:10.1073/pnas.0510565103
Wada, T., Kikuchi, J., and Furukawa, Y. (2012). Histone deacetylase 1 enhances microRNA processing via deacetylation of DGCR8. EMBO Rep. 13, 142–149. doi:10.1038/embor.2011.247
Wagner, S. A., Beli, P., Weinert, B. T., Nielsen, M. L., Cox, J., Mann, M., et al. (2011). A proteome-wide, quantitative survey of in vivo ubiquitylation sites reveals widespread regulatory roles. Mol. Cell. Proteomics MCP 10, M111.013284. doi:10.1074/mcp.M111.013284
Walz, A. L., Ooms, A., Gadd, S., Gerhard, D. S., Smith, M. A., Guidry Auvil, J. M., et al. (2015). Recurrent DGCR8, DROSHA, and SIX homeodomain mutations in favorable histology wilms tumors. Cancer Cell 27, 286–297. doi:10.1016/j.ccell.2015.01.003
Wang, W., and Matunis, M. J. (2023). Paralogue-specific roles of SUMO1 and SUMO2/3 in protein quality control and associated diseases. Cells 13, 8. doi:10.3390/cells13010008
Wang, Y., Medvid, R., Melton, C., Jaenisch, R., and Blelloch, R. (2007). DGCR8 is essential for microRNA biogenesis and silencing of embryonic stem cell self-renewal. Nat. Genet. 39, 380–385. doi:10.1038/ng1969
Wang, G., Gao, Y., Li, L., Jin, G., Cai, Z., Chao, J.-I., et al. (2012). K63-linked ubiquitination in kinase activation and cancer. Front. Oncol. 2, 5. doi:10.3389/fonc.2012.00005
Weber, J. D., Taylor, L. J., Roussel, M. F., Sherr, C. J., and Bar-Sagi, D. (1999). Nucleolar Arf sequesters Mdm2 and activates p53. Nat. Cell Biol. 1, 20–26. doi:10.1038/8991
Weber, C., Schreiber, T. B., and Daub, H. (2012). Dual phosphoproteomics and chemical proteomics analysis of erlotinib and gefitinib interference in acute myeloid leukemia cells. J. Proteomics 75, 1343–1356. doi:10.1016/j.jprot.2011.11.004
Wei, B., Yang, F., Yu, L., and Qiu, C. (2024). Crosstalk between SUMOylation and other post-translational modifications in breast cancer. Cell. Mol. Biol. Lett. 29, 107. doi:10.1186/s11658-024-00624-3
Wostenberg, C., Quarles, K. A., and Showalter, S. A. (2010). Dynamic origins of differential RNA binding function in two dsRBDs from the miRNA “microprocessor” complex. Biochemistry 49, 10728–10736. doi:10.1021/bi1015716
Xia, C., Tao, Y., Li, M., Che, T., and Qu, J. (2020). Protein acetylation and deacetylation: an important regulatory modification in gene transcription (Review). Exp. Ther. Med. 20, 2923–2940. doi:10.3892/etm.2020.9073
Yang, Q., Li, W., She, H., Dou, J., Duong, D. M., Du, Y., et al. (2015). Stress induces p38 MAPK-mediated phosphorylation and inhibition of DROSHA-dependent cell survival. Mol. Cell 57, 721–734. doi:10.1016/j.molcel.2015.01.004
Ye, P., Liu, Y., Chen, C., Tang, F., Wu, Q., Wang, X., et al. (2015). An mTORC1-Mdm2-DROSHA axis for miRNA biogenesis in response to glucose- and amino acid-deprivation. Mol. Cell 57, 708–720. doi:10.1016/j.molcel.2014.12.034
Yin, S., Yu, Y., and Reed, R. (2015). Primary microRNA processing is functionally coupled to RNAP II transcription in vitro. Sci. Rep. 5, 11992. doi:10.1038/srep11992
Zhou, H., Di Palma, S., Preisinger, C., Peng, M., Polat, A. N., Heck, A. J. R., et al. (2013). Toward a comprehensive characterization of a human cancer cell phosphoproteome. J. Proteome Res. 12, 260–271. doi:10.1021/pr300630k
Zhu, C., Chen, C., Huang, J., Zhang, H., Zhao, X., Deng, R., et al. (2015). SUMOylation at K707 of DGCR8 controls direct function of primary microRNA. Nucleic Acids Res. 43, 7945–7960. doi:10.1093/nar/gkv741
Keywords: microprocessor, DROSHA, DGCR8, microRNA biogenesis, phosphorylation, acetylation, ubiquitination, SUMOylation
Citation: Leong KW and Chong MMW (2025) Regulation of the microprocessor by post-translational modifications. Front. Cell Dev. Biol. 13:1721087. doi: 10.3389/fcell.2025.1721087
Received: 09 October 2025; Accepted: 19 November 2025;
Published: 01 December 2025.
Edited by:
Congbao Kang, Experimental Drug Development Centre (EDDC), SingaporeReviewed by:
Hejer Dhahri, George Washington University, United StatesCopyright © 2025 Leong and Chong. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Mark M. W. Chong, bWNob25nQHN2aS5lZHUuYXU=