D-type cyclins (cyclin D1, cyclin D2, and cyclin D3) are activators of the cyclin-dependent kinases CDK4 and CDK6 and represent major oncogenic drivers among members of the cyclin superfamily (Musgrove et al., 2011; Fassl et al., 2022; Álvarez-Fernández and Malumbres, 2020; Behan et al., 2019). The CCND1 gene --encoding cyclin D1-- shows some of the highest frequency of amplification and overexpression among cancer genes across a broad spectrum of human tumors (Beroukhim et al., 2010; Curtin et al., 2005; Bignell et al., 2010; Parsons et al., 2008; Jones et al., 2008). Moreover, mutations in CCND1, which prevent the degradation of cyclin D1 by AMBRA1, a substrate receptor (SR) of a CUL4-RING ubiquitin ligase (CRL4) complex that targets all three D-type cyclins for proteasome-mediated degradation, have been reported in a variety of tumor types (Simoneschi et al., 2021; Chaikovsky et al., 2021; Martínez-Jiménez et al., 2020). Deregulation of cellular proliferation, often mediated by uncontrolled CDKs’ activation lies at the heart of cancer as a pathological process (Kastan and Bartek, 2004).

The biological roles of D-type cyclins have been examined almost exclusively through the lens of E2F transcription regulation upon phosphorylation of the three pocket proteins, RB, p107, and p130. D-type cyclin-mediated phosphorylation of RB results in the inactivation of its tumor suppressive effect by releasing E2F from the RB’s inhibitory effect on gene transcription. Although the regulation of RB by phosphorylation is well understood in both physiology and human tumorigenesis, the cancer-related role of phosphorylation of other substrates is a subject that has remained understudied. In this study, we characterize GTSE1 (G-Two and S phase expressed protein 1) as a previously unidentified substrate of cyclin D1–CDK4/6 in cells overexpressing cyclin D1 in G1 and beyond. GTSE1 is a cell cycle-related protein expressed specifically during the S and G2 phases of the cell cycle (Liu et al., 2010). It interacts with the tumor suppressor p53, and it induces its MDM2-mediated degradation in G2 and during the recovery from DNA damage, promoting cell proliferation (Liu et al., 2010; Bublik et al., 2010; Monte et al., 2003; Monte et al., 2004; Collavin et al., 2000; Utrera et al., 1998). Moreover, GTSE1 associates with growing microtubules, promoting cell migration (Bendre et al., 2016; Scolz et al., 2012). In prometaphase, GTSE1 becomes highly phosphorylated by CDK1–cyclin B1, resulting in its recruitment to the inner spindle (Singh et al., 2021). After anaphase, its dephosphorylation is followed by marked reduction in its abundance in G1 (Collavin et al., 2000; Monte et al., 2000). Our findings demonstrate that the cyclin D1–CDK4/6-mediated phosphorylation of GTSE1 leads to its increased stability in G1 phase, an event that significantly impacts cell proliferation and cancer prognosis.

AMBRA1 is an SR of a CUL4-RING ubiquitin ligase (CRL4) complex that targets all three D-type cyclins for proteasome-mediated degradation (Simoneschi et al., 2021; Chaikovsky et al., 2021). To unveil new substrates of the cyclin D1–CDK4 complex, we performed a comprehensive multi-analysis of published data (Figure 1—figure supplement 1A). First, we incorporated recent mass spectrometry data that compared the whole proteome of AMBRA1 knockout (KO) clones to parental U2OS cells (Chaikovsky et al., 2021), aiming to pinpoint the top 30 proteins whose levels were elevated in the absence of AMBRA1, the SR of the ubiquitin ligase targeting D-type cyclins (Figure 1A). Subsequently, we asked which of these 30 proteins contain a canonical CDK phosphorylation consensus motif [S/T*]PX[K/R], relying on the PhosphoSitePlus database (Hornbeck et al., 2015; Figure 1B). Finally, this dataset was integrated with findings from a proteomic screen assessing protein abundance fluctuations in the presence or absence of the CDK4/6 inhibitor, Palbociclib (Figure 1C; Chaikovsky et al., 2021). In summary, our analysis aimed at identifying proteins whose upregulation in AMBRA1 KO cells was counteracted by Palbociclib treatment, thereby filtering for proteins whose augmented abundance is attributed to CDK4/6-mediated phosphorylation events when cyclin D1 is stabilized. GTSE1, a cell cycle-regulated protein expressed mainly during the G2 and S phases of the cell cycle (Liu et al., 2010), emerged as the top hit in these orthogonal analyses. In mitosis GTSE1 is phosphorylated by CDK1–cyclin B1 (Singh et al., 2021), but its dephosphorylation at the end of mitosis is followed by marked reduction in its abundance, which remains low during the next G1 (Collavin et al., 2000; Monte et al., 2000).

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Recent pan-cancer analysis revealed that the expression of GTSE1 positively correlates with tumor mutational burden and microsatellite instability in most cancer types (Tan et al., 2023). Specifically, high expression of GTSE1 was found to promote the proliferation and invasion of breast cancer cells (Lin et al., 2019), and was associated with poor clinical prognosis in clear cell renal cell carcinoma (Lei et al., 2023). We explored further the potential influence of GTSE1 on cancer prognosis and found that within multiple cancer cohorts, its elevated expression levels were correlated with a statistically significant poorer prognosis compared to lower expression levels (Figure 1D, Figure 1—figure supplement 1B). This is noteworthy, since, except for acute myeloid leukemia (LAML), GTSE1 expression was higher than that in normal tissue counterparts in all tumor types analyzed (Figure 1E). We also assessed the impact of GTSE1 on survival across various cancer types (Figure 1F, Figure 1—figure supplement 1B). For context, we compared the survival patterns associated with cyclin D1, CDK4, and CDK6. Similar to the overexpression of other cell cycle-regulated genes (Whitfield et al., 2006), high levels of GTSE1 were found to be prognostically unfavorable across multiple cancer types, presenting a survival pattern similar to that of the cyclin D1–CDK4/6 gene cluster. In Figure 1F, those cancers in which differences in survival are statistically significant were denoted by contour squares.

Next, we sought to experimentally validate GTSE1 as a putative phosphorylation target of cyclin D1–CDK4. As a first step, we employed transient transfection to introduce in HEK293T various Flag-tagged constructs in the presence or absence of cyclin D1 and CDK4. GTSE1 showed an upper shift in a phos-tag gel when co-expressed with cyclin D1–CDK4, similar to the known substrates p107 and p130 (Figure 2A). In fact, a slight delay in GTSE1 migration is appreciable even in a regular SDS–PAGE (see first two lanes in Figure 2A). The upper shift was not observed in ZC3HAV1, a protein used as negative control. Next, we performed an in vitro phosphorylation assay to directly assess the specificity of GTSE1 phosphorylation by cyclin D1–CDK4. Using purified, recombinant proteins, we subjected GTSE1 to a phosphorylation reaction with cyclin D1–CDK4 and analyzed the products using phos-tag gels. The results confirmed that GTSE1 underwent phosphorylation by cyclin D1–CDK4, but not by ERK1, another Pro-directed kinase (Figure 2B). Moreover, the phosphorylation was abolished in the presence of Palbociclib, similar to the pattern observed with RB (Figure 2B, C, H). Then, we asked whether GTSE1 physically interacts with the kinase complex. Overexpression of Flag-tagged cyclin D1 and CDK4 in HEK293T followed by a Flag pull-down demonstrated binding to endogenous GTSE1, similar to the canonical substrate RB (Figure 2D).

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Original files for western blot analysis displayed in Figure 2.

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PDF file containing original western blots for Figure 2, indicating the relevant bands.

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The raw data for the box plot in Figure 2I depicting GTSE1 phosphorylation levels normalized to total GTSE1 protein abundance across a pan-cancer cohort is provided as log2(Intensities) across samples for global proteins and phospho-proteins of interest.

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The pre-processed data for the box plot in Figure 2I and the heatmap illustrating differential abundance of GTSE1 phosphorylation sites in Figure 2J is provided as Row Z-score of GTSE1 log2(Intensities) across samples for global protein and phospho-protein.

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Following these validation steps, we aimed at pinpointing the serine residue(s) modified by cyclin D1–CDK4. First, we generated a GTSE1 mutant in which Ser262 (see Figure 1B) was mutated to Ala. Since GTSE1 has additional, conserved serine residues followed by prolines, which could be potential CDK phosphorylation sites (Figure 2E, Figure 2—figure supplement 1A, B), we also generated another sixteen Ser-to-Ala GTSE1 mutants and expressed them in HEK293T cell in the presence or absence of cyclin D1–CDK4 (Figure 3—figure supplement 1A). When compared to wild-type GTSE1, two mutants displayed changes in their migrations on phos-tag gels (Figure 3—figure supplement 1A), suggesting that Ser residues at positions 91 and 724 are sites potentially phosphorylated by cyclin D1–CDK4. PhosphoSitePlus (Hornbeck et al., 2015) reports several high-throughput studies that, in addition to Ser262, identified phosphorylation also at Ser91 and Ser724, supporting our results. To corroborate the loss of cyclin D1–CDK4-dependent phosphorylation at these sites, we constructed a triple mutant (S91A/S262A/S724A). This triple mutant displayed loss of slower-migrating bands relative to wild-type GTSE1, suggesting diminished phosphorylation (Figure 2F). Nevertheless, a residual slow-migrating band persisted, prompting further mutations of the triple GTSE1 mutant in two additional GTSE1 sites (individually), which do not have a CDK-phosphorylation consensus, but were identified in several proteomics studies (Hornbeck et al., 2015; Kaulich et al., 2021). From these two mutants, only the S454A mutation demonstrated a complete abrogation of any shift in phos-tag gels (Figure 2F). In contrast, a single S454A mutation showed only a minor effect on GTSE1 shift (data not shown). These studies suggest that four major sites (S91, S262, S454, and S724) are phosphorylated (either directly and/or indirectly) in a cyclin D1–CDK4-dependent manner.

To gain an insight into the cell cycle pattern of GTSE1 abundance and phosphorylation status, we synchronized T98G cells (both parental and AMBRA1 KO pooled clones) by serum starvation followed by serum re-addition. Parental cells displayed a fluctuation in GTSE1 abundance, with low levels in G1 that subsequently increased in S and G2 phases (Figure 3—figure supplement 1B), in agreement with the literature (Collavin et al., 2000; Monte et al., 2000). In contrast, in AMBRA1 KO cells, GTSE1 levels were consistently elevated throughout the cell cycle, suggesting that elevated cyclin D1 levels are associated with an increased abundance of GTSE1 (Figure 3—figure supplement 1B). Additionally, while phosphorylation of GTSE1 in the parental cell line peaked during the G2/M transition (28–32 hr after serum re-addition), AMBRA1 KO cells exhibited sustained phosphorylation of GTSE1 across all cell cycle phases (Figure 2G, Figure 3—figure supplement 1B). We also generated a phospho-specific antibody that recognized GTSE1 only when phosphorylated on Ser262 (Figure 3—figure supplement 1C). Using this antibody, we confirmed the data obtained with phos-tag gel indicating that in cells overexpressing cyclin D1–CDK4 and in AMBRA1 KO cells, GTSE1 is hyper-phosphorylated on Ser262 (Figure 2G, Figure 3—figure supplement 1C).

Next, we leveraged data from the Clinical Proteomic Tumor Analysis Consortium (CPTAC) to examine the relevance of the identified phosphorylation events in a clinical context. We observed an enrichment of GTSE1 phospho-peptides within a pan-cancer cohort as opposed to adjacent, corresponding normal tissues (Figure 2I), underscoring the potential role of GTSE1 phosphorylation in tumorigenesis. Upon analyzing single phosphorylated sites, we found that nine were statistically enriched in cancers. Of these, three sites (S262, S454, and S724) were identified as dependent on cyclin D1–CDK4 in our study (Figure 2J, Figure 3—figure supplement 1D). (Data on Ser91 are not present in CPTAC.) These results underscore the potential pathophysiological significance of GTSE1 phosphorylation by cyclin D1–CDK4 in cancer.

Considering GTSE1 being an established target of cyclin B1–CDK1 during mitosis (Singh et al., 2021), we aimed to elucidate the phosphorylation patterns of GTSE1 by various cyclin–CDK complexes operating at distinct cell cycle phases. Wild-type GTSE1 was found to be phosphorylated in HEK293T cells upon overexpression of all cyclin–CDK pairs, with its shifts in phos-tag gels being abolished upon treatment with the corresponding specific CDK inhibitor (Figure 2H, left panel). However, when the quadruple (S91A/S262A/S454A/S724A) GTSE1 mutant, referred to as ‘Tetra SA’, was expressed in HEK293T cells, cyclin D1–CDK4 was unable to induce any shift, whereas the other three cyclin–CDK pairs still sustained phosphorylation (Figure 2H, right panel). This suggests a unique phosphorylation profile conferred by cyclin D1–CDK4 in GTSE1, distinct from that induced by other cyclin–CDK complexes.

We noticed an elevation in endogenous GTSE1 levels in AMBRA1 KO cells compared to the parental line (Figure 3A, Figure 3—figure supplement 1B) in agreement with the findings of the proteomic screen performed in AMBRA1 KO cells (Chaikovsky et al., 2021; Figure 1B). A similar increase was observed when overexpressing wild-type cyclin D1 and, even more, an AMBRA1-insensitive, stable mutant of cyclin D1 (T286A) (Simoneschi et al., 2021; Figure 3A, B), suggesting that the elevated GTSE1 levels are due to high D-type cyclins present in AMBRA1 KO cells. We also used HCT-116 cells harboring an endogenous fusion of AMBRA1 to a minimally constructed Auxin Inducible Degron (mAID) at the N-terminus (Yesbolatova et al., 2020). This system allows for rapid and inducible degradation of AMBRA1 upon addition of auxin, thereby minimizing compensatory cellular rewiring. Again, we observed an increase in GTSE1 levels upon acute ablation of AMBRA1 (i.e., in 8 hr) (Figure 3B). In all cases, the upregulation in GTSE1 abundance was rescued upon Palbociclib treatment (Figure 3A, B), suggesting that this event was a consequence of increased levels of D-type cyclins. We also conducted cycloheximide (CHX) chase assays in HCT-116 mAID-AMBRA1 cells to assess GTSE1 protein stability and degradation kinetics. The assays revealed that in the context of AMBRA1 depletion, GTSE1 exhibited a prolonged half-life and reduced degradation rate when compared to control cells (Figure 3C). A parallel half-life assessment in parental U2OS cells and two AMBRA1 KO U2OS clones corroborated the finding that GTSE1 is stabilized in the absence of AMBRA1 (Figure 3D).

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To further dissect the impact of cyclin D1–CDK4-mediated phosphorylation on GTSE1 stability, we engineered a phospho-mimicking mutant, referred to as ‘Tetra SD’ with the four serine residues replaced with an aspartate at positions 91, 261, 454, and 724. To circumvent the variability of transient transfection, U2OS cells were stably transduced with retroviruses encoding GFP-tagged wild-type GTSE1, Tetra SA (phospho-deficient), or Tetra SD (phospho-mimic). Subsequent CHX chase experiments showed a slower degradation kinetics of the Tetra SD mutant compared to the Tetra SA mutant and wild-type GTSE1 (Figure 3E, F). These stable cell lines, expressing fluorescent GTSE1 variants, were further analyzed via time-lapse microscopy during a CHX chase to quantify protein degradation through the diminishing fluorescence intensity at different times. The Tetra SD mutant exhibited a statistically significant slower fluorescence decrease rate than both Tetra SA and WT, indicating reduced degradation (Figure 3G, H).

Finally, we treated U2OS cells with CHX together with inhibitors targeting different degradation systems: MG132 (a proteasome inhibitor), MLN4924 (a CRL inhibitor), or Bafilomycin A (an autophagy inhibitor) (Figure 3I). The stabilization of GTSE1 in the presence of MG132 and MLN4924, but not Bafilomycin A, indicates that GTSE1 degradation mainly occurs via the ubiquitin–proteasome system, specifically implicating the involvement of a CRL. However, GTSE1 does not appear to be a substrate of CRL4^AMBRA1, as indicated by the lack of a physical interaction between AMBRA1 and GTSE1 (Figure 3—figure supplement 1E), as well as the fact that the stabilization of GTSE1 in AMBRA1 KO cells is secondary to the CDK4/6 activity (Figure 3A, B).

Next, we delved into the potential implications of GTSE1 phosphorylation by the cyclin D1–CDK4 complex. Previous research has connected GTSE1 with fundamental cellular processes, such as cell proliferation (Lei et al., 2023; Lai et al., 2021) and cell migration and invasion (Scolz et al., 2012; Wu et al., 2017; Li et al., 2021). Leveraging data from CPTAC, we examined possible correlations between both GTSE1 protein levels (Figure 4—figure supplement 1A) and levels of phosphorylated GTSE1 (Figure 4A) with markers of cell proliferation and cell migration across various cancer types. GTSE1 abundance and phosphorylation exhibited a statistically significant positive correlation with several proteins integral to cell proliferation, including PCNA, KI67, and MCM2, across multiple cancer cohorts. Conversely, in contrast to data suggesting a role for GTSE1 in promoting cell migration (Scolz et al., 2012; Lin et al., 2019; Lei et al., 2023; Wu et al., 2017) GTSE1 levels and phosphorylation demonstrated a negative trend with proteins associated with cell migration and epithelial–mesenchymal transition, such as Vimentin, MMP1, and ETS1, but these did not attain statistical significance.

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The source data for Figure 4A is provided as Pairwise GTSE1 (phospho) ~ Hallmark (protein) Spearman correlations from proteomic input.

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To explore the influence of cyclin D1-mediated phosphorylation of GTSE1 on cellular phenotypes, we assessed the proliferation potential of AMBRA1 KO cells relative to their parental counterparts. AMBRA1 KO cells displayed an enhanced proliferation rate (Figure 4—figure supplement 1B), in agreement with the literature (Simoneschi et al., 2021). We then conducted comparable growth analyses using U2OS cells stably expressing various GTSE1 constructs. Cells stably expressing the Tetra SD mutant exhibited a higher proliferation rate compared to cells expressing either Tetra SA or WT GTSE1 (Figure 4B). Moreover, evaluations of cellular proliferation conducted using Cell Trace dye demonstrated that cells harboring the Tetra-SD mutant displayed an elevated proliferative index (Figure 4C, Figure 4—figure supplement 1D, E). This was evidenced by a greater number of cell generations and smaller fraction of non-divided cells relative to cells expressing WT GTSE1 and, even more so compared to cells expressing Tetra SA (Figure 4C, Figure 4—figure supplement 1D, E). Finally, cell cycle profiles showed that the expression of a phospho-mimicking GTSE1 mutant induces a decrease in the percentage of cells in G1 and an increase in the percentage of cells in S, similarly to what observed in AMBRA1 KO cells (Figure 4—figure supplement 1F and Rona et al., 2024).

This study primarily analyzed publicly available datasets from the Clinical Proteomic Tumor Analysis Consortium (CPTAC). The CPTAC datasets used in this study include pan-cancer proteomic data referenced for the analysis of GTSE1 phosphorylation levels across multiple cancer types (Figure 2I and J, Figure 4A, Figure 3—figure supplement 1D, and Figure 4—figure supplement 1A). All analytical results generated during this study are included in the manuscript and supporting files. Source data files have been provided for the relevant figures. Uncropped gel scans used for figure generation have been deposited in Mendeley Data Repository (https://doi.org/10.17632/xzkw7hrwjr.1). Plasmids generated in this study have been deposited in Addgene. Cell lines and the phospho-specific antibody against GTSE1 phosphorylated on Ser262 generated in this study are available from the corresponding authors upon reasonable request. All other materials, reagents, and computational tools described in this paper are available from the authors or from standard commercial sources as specified in the Materials and methods section. CPTAC data access: The CPTAC datasets used in this study are available through the CPTAC Data Portal (https://pdc.cancer.gov/pdc/cptac-pancancer) and the Genomic Data Commons (https://portal.gdc.cancer.gov/). Specifically, we used the pre-processed CPTAC datasets Proteome_UMich_SinaiPreprocessed_GENECODE34_v1.zip and Phosphoproteome_UMich_SinaiPreprocessed_GENECODE34_v1.zip. Access to the raw datasets requires approval through the dbGaP authorization process (https://dbgap.ncbi.nlm.nih.gov/), in accordance with the data access policies established by the National Cancer Institute. CPTAC data citation: In accordance with CPTAC data usage policies, we have cited the appropriate CPTAC publications and acknowledged the CPTAC consortium as the source of primary data. Specifically, we used the following CPTAC datasets: Pan-cancer proteomic and phosphoproteomic data as referenced in Ellis et al., 2013, and The Cancer Genome Atlas (TCGA) data for transcriptomic analysis as referenced in The Cancer Genome Atlas Research Network et al., 2013. The proteomic data from AMBRA1 knockout U2OS cells compared to parental U2OS cells referenced in Figure 1A–C was obtained from previously published work by Chaikovsky et al., 2021. This dataset, titled "Shotgun mass spectrometry analysis comparing AMBRA1 KO U2OS cell populations and wild-type controls," is available in the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD021789.

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Author details

1. Nelson García-Vázquez

   Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States

   Contribution

   Formal analysis, Investigation

   Competing interests

   No competing interests declared

2. Tania J González-Robles

   1. Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States
   2. Department of Medicine, New York University Grossman School of Medicine, New York, United States
   3. Howard Hughes Medical Institute, New York University Grossman School of Medicine, New York, United States

   Contribution

   Data curation, Visualization

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-9292-382X

3. Ethan Lane

   Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States

   Contribution

   Investigation

   Competing interests

   No competing interests declared

4. Daria Spasskaya

   Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States

   Contribution

   Investigation

   Competing interests

   No competing interests declared

5. Qingyue Zhang

   Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States

   Contribution

   Investigation

   Competing interests

   No competing interests declared

6. Marc A Kerzhnerman

   Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States

   Contribution

   Investigation

   Competing interests

   No competing interests declared

7. YeonTae Jeong

   Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States

   Present address

   Arvinas Inc, New Haven, United States

   Contribution

   Investigation

   Competing interests

   is currently an employee of Arvinas, Inc, contributions to this paper were completed before joining Arvinas

8. Marta Collu

   Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States

   Contribution

   Investigation

   Competing interests

   No competing interests declared

9. Daniele Simoneschi

   Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States

   Contribution

   Investigation

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-0448-4295

10. Kelly V Ruggles

    Department of Medicine, New York University Grossman School of Medicine, New York, United States

    Contribution

    Resources, Visualization

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-0152-0863

11. Gergely Róna

    1. Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States
    2. Howard Hughes Medical Institute, New York University Grossman School of Medicine, New York, United States
    3. Institute of Molecular Life Sciences, HUN-REN Research Centre for Natural Sciences, Budapest, Hungary

    Contribution

    Investigation

    Competing interests

    No competing interests declared

12. Sharon Kaisari

    1. Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States
    2. Howard Hughes Medical Institute, New York University Grossman School of Medicine, New York, United States

    Contribution

    Conceptualization, Supervision, Investigation, Writing – original draft, Writing – review and editing

    For correspondence

    sharon.kaisari@nyulangone.org

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-6884-3886

13. Michele Pagano

    1. Department of Biochemistry and Molecular Pharmacology, New York University Grossman School of Medicine, New York, United States
    2. Howard Hughes Medical Institute, New York University Grossman School of Medicine, New York, United States

    Contribution

    Conceptualization, Funding acquisition, Writing – original draft, Writing – review and editing

    For correspondence

    michele.pagano@nyulangone.org

    Competing interests

    is or has been an advisor for and has financial interests in SEED Therapeutics, Triana Biomedicines, CullGen, Kymera Therapeutics, Serinus Biosciences, and Umbra Therapeutics

    "This ORCID iD identifies the author of this article:" 0000-0003-3210-2442

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