Abstract

Microcephaly primary hereditary (MCPH) is a rare, neurological disorder characterized by a small brain size, due to a lower number of neural progenitor cells, and mental retardation (Naveed in Genetics Research 100:e7, 2018). Approximately 40% of all MCPH patients harbor mutations in abnormal spindle-like microcephaly-associated (ASPM) gene, which is also known as MCPH5 (Létard in Human Mutation 39(3):319–32, 2018). The ASPM protein is involved in centriole duplication, orientation of the spindle, and regulation of mitosis (Jiang in Nature Cell Biology 19(5):480–492, 2017; Gai in EMBO Reports 17(10):1396–1409, 2016). Although these functions of ASPM seem to be independent of DNA replication, impaired DNA replication has been associated with microcephaly (Tingler in Biology of the Cell 114(6):143–159; Kalogeropoulou in Stem Cell Reports 17(6):1395–1410; Xu in Genome Instability & Disease 1:235–264). In a recent PNAS paper by Wu et al., the authors suggested that patients with mutations in ASPM harbor a reduced number of neuroprogenitor cells due to defects in the DNA stress response.

Defective replication stress response linked to microcephaly

To identify functions of ASPM during DNA replication, Wu et al. generated a knock-in cell line by inserting a Flag–GFP cassette in frame before the initiation codon of the endogenous ASPM gene, thereby expressing endogenous ASPM as a Flag–GFP–ASPM fusion protein. First, they sought to determine the interacting partners of ASPM and for that, they performed immunoprecipitation of the endogenous Flag–GFP tagged ASPM followed by mass spectrometry. Using this approach, they identified that ASPM directly interacts with several proteins involved in DNA replication, including subunits of the replication factor C (RFC) complex (RFC1-5), TopBP1, and RAD17. Since ASPM interacts with DNA replication factors, Wu et al. generated ASPM knockout HeLa cells and tested whether loss of ASPM has a direct impact on DNA replication. They observed that loss of ASPM had no particular impact on the rate of replication fork progression or on the progression of cells through S-phase, suggesting that ASPM does not affect replication under unperturbed conditions.

Nucleotide depletion, polymerase inhibition or DNA damage cause replication fork stalling and induce replication stress (Zeman & Cimprich, 2014). Replication stress in turn leads to the phosphorylation and activation of the ATR and CHK1 kinases, which mediate the replication stress response and promote recovery of stalled forks by stabilizing them and allowing them to restart (Hsiao et al., 2021); Liu et al., 2011; Zou & Elledge, 2003). Interestingly, Wu et al. found that ASPM is required for the activation of ATR and CHK1 upon DNA replication stress induction by hydroxyurea (HU), aphidicolin (APH), or camptothecin (CPT). ASPM KO cells had a defect in the phosphorylation of ATR at T1989 and CHK1 at S345. In addition, they observed that ASPM loss leads to an arrest of cells in S-phase upon HU or APH treatment. Moreover, ASPM KO cells exhibited a reduction in phosphorylated RPA (pRPA) foci and native BrdU staining upon treatment with HU, indicating a defect in the formation of single-stranded DNA (ssDNA).

Wu et al. then sought to study the functional importance of the interaction between ASPM and the RFC complex, RAD17, and TopBP1. RAD17 promotes loading of RAD9–RAD1–HUS1 (9–1–1) complexes and of TopBP1 onto stalled replication forks, thereby promoting downstream activation of the ATR–CHK1 axis (Zou et al., 2002, 2003). Using immunoprecipitation, they found that the interaction of ASPM with RAD17, RAD9, and TopBP1 was enhanced upon replication stress induction by HU treatment. Surprisingly, upon HU treatment, loss of ASPM reduced the chromatin levels of RAD9, TopBP1, and RPA, but not of RAD17, suggesting that RAD17 is upstream of ASPM in recruiting 9–1–1 complex and TopBP1 at stalled forks. Consistent with this idea, loss of RAD17 reduced the chromatin binding of RAD9, TopBP1, and ASPM upon HU treatment. Moreover, the reduced induction of pRPA by HU in ASPM KO cells could be rescued by over-expressing RAD9 or the ATR activation domain of TopBP1 (TopBP1–AAD), but not RAD17. Together, the results of Wu et al. suggest that ASPM aids the RAD17-dependent loading of 9–1–1 and TopBP1 onto chromatin upon replication stress.

Using the isolation of Proteins On Nascent DNA (iPOND) method (Sirbu et al., 2011), the authors found that ASPM was enriched onto nascent DNA upon HU treatment. They performed proximity ligation assays (PLA) to detect the colocalization of EdU-labeled nascent DNA with RAD17, RAD9, and TopBP1, to test whether the recruitment of RAD9, TopBP1, and RAD17 to replication forks is dependent on ASPM. They found no change in the RAD17–EdU PLA foci in the presence or absence of ASPM, whereas they saw a clear reduction in RAD9–EdU and TopBP1–EdU PLA foci. These findings were further confirmed using in vitro streptavidin pulldown assays in which the authors used RPA-coated biotinylated single-stranded/double-stranded DNA substrates to pulldown proteins from nuclear extracts of WT or ASPM KO cells. Interestingly, RPA-coated biotinylated ss/dsDNA substrates only pulled down RAD17 from ASPM KO extracts but not RAD9 and TopBP1. However, RPA-coated ss/dsDNA substrates saturated with RAD17 were able to pull down RAD9 and TopBP1 only in the presence of ASPM. These results suggest that ASPM promotes the RAD17-dependent loading of 9–1–1 onto stalled forks upon replication stress.

Since ASPM localizes to newly synthesized DNA at stalled replication forks, Wu et al. examined whether ASPM is involved in fork protection and found that ASPM does protect reversed forks against MRE11-mediated but not EXO1- or DNA2-mediated nucleolytic degradation. Based on this finding, the authors speculated that loss of ASPM should lead to an overall increase in genomic instability under conditions of replication stress. Indeed, metaphase spreads performed to examine gross chromosomal defects showed higher levels of chromosome breaks, fusions, and other abnormalities in the absence of ASPM and upon replication stress. Interestingly, the authors also report that high ASPM expression in several types of cancer associates with an overall poorer prognosis, suggesting that ASPM inhibition could be a therapeutic strategy targeting tumors dependent on high ASPM levels (Feng et al., 2021; Xu et al., 2019).

While this study shows that ASPM is involved in DNA replication by protecting stalled forks, several questions remain. Recently, the same group has shown that ASPM promotes BRCA1 stability by preventing degradation of BRCA1 by HERC2 and that loss of ASPM compromises homologous recombination (HR) (Xu et al., 2021). Moreover, ASPM loss also leads to increased sensitivity to PARP inhibition (Xu et al., 2021). These phenotypes observed upon loss of ASPM almost resemble ‘BRCAness’, a condition which phenocopies loss of BRCA1/2 and is manifested by defects in HR, increased genomic instability, replication fork protection defects, and sensitivity to PARP inhibitors (Byrum et al., 2019). Are the ‘BRCAness’ phenotypes shown by the loss of ASPM direct or indirect? Specifically with respect to this study, it would be interesting to investigate whether the fork protection functions of ASPM are dependent on either BRCA1 or BRCA2 since ASPM was also found to interact with both BRCA1 and BRCA2 (Xu et al., 2021). It is likely that in the absence of ASPM, BRCA1 in association with its obligate partner BARD1, or BRCA2 can no longer protect against degradation of the nascent strand (Daza-Martin et al., 2019; Schlacher et al., 2011; Taglialatela et al., 2017). Another question is how ASPM gets recruited to stalled forks. Using immunofluorescent microscopy methods, it was previously observed that ASPM is recruited in a PARP2-dependent manner at sites of DNA breaks where it promotes DNA end resection (Xu et al., 2021). Is ASPM recruited to stalled forks through PARP2 as well? Does RAD17 function independently of PARP2 or together with PARP2 to recruit ASPM?

In conclusion, ASPM facilitates the replication stress response by not only promoting the activation of the ATR–CHK1 axis via the loading of 9–1–1 and TopBP1, but also by protecting nascent strands from degradation. The addition of ASPM to the ever-expanding and scopious arsenal of proteins that sense DNA replication stress and mount efficient cellular responses simply highlights the importance of protecting DNA during synthesis to prevent human developmental diseases.

Data availability statement

No original data is generated in this study.

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Ackonwledgements

Lee Zou is the James and Patricia Poistra Endowed Chair of Cancer Reserach.

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Authors and Affiliations

1. Massachusetts General Hospital Cancer Center, Harvard Medical School, Charlestown, MA, 02129, USA

   Ajinkya S. Kawale & Lee Zou

2. Department of Pathology, Harvard Medical School, Massachusetts General Hospital, Boston, MA, 02114, USA

   Lee Zou

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Correspondence to Lee Zou.

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Kawale, A.S., Zou, L. Defective replication stress response linked to microcephaly. GENOME INSTAB. DIS. 3, 267–269 (2022). https://doi.org/10.1007/s42764-022-00084-z

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• Received10 September 2022

• Revised10 September 2022

• Accepted18 September 2022

• Published11 October 2022

• Issue DateOctober 2022

• DOIhttps://doi.org/10.1007/s42764-022-00084-z

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Keywords

• Microcephaly

• Replication stress

• ASPM

• ATR

• TopBP1