PPP1R35 is a novel centrosomal protein that regulates centriole length in concert with the microcephaly protein RTTN
Abstract
Centrosome structure, function, and number are finely regulated at the cellular level to ensure normal mammalian development. Here, we characterize PPP1R35 as a novel bona fide centrosomal protein and demonstrate that it is critical for centriole elongation. Using quantitative super-resolution microscopy mapping and live-cell imaging we show that PPP1R35 is a resident centrosomal protein located in the proximal lumen above the cartwheel, a region of the centriole that has eluded detailed characterization. Loss of PPP1R35 function results in decreased centrosome number and shortened centrioles that lack centriolar distal and microtubule wall associated proteins required for centriole elongation. We further demonstrate that PPP1R35 acts downstream of, and forms a complex with, RTTN, a microcephaly protein required for distal centriole elongation. Altogether, our study identifies a novel step in the centriole elongation pathway centered on PPP1R35 and elucidates downstream partners of the microcephaly protein RTTN.
https://doi.org/10.7554/eLife.37846.001eLife digest
Most animal cells contain a structure called the centrosome, which plays a vital role in helping cells to divide for producing new cells. Early in the cell division process, cells make a copy of their centrosome. Each centrosome includes two cylindrical structures called centrioles encased in a complex web of other proteins. The centrioles must get longer for the duplication process to work correctly, but it is not clear which proteins help the centrioles to elongate.
Previous work suggested that a protein called PPP1R35 might be a centrosome protein. To investigate its role, Sydor et al. performed experiments that reduced the amount of PPP1R35 in cells grown in the laboratory. Cells that contained fewer PPP1R35 proteins also contained fewer centrioles; these centrioles were also shorter and lacked some of the proteins that can elongate them.
Super-resolution microscopy found PPP1R35 in the centre of the centrioles, in a region involved in the early stages of elongation. Sydor et al. also found that PPP1R35 interacts with a protein called RTTN, which is linked to centriole elongation.
RTTN contributes to a condition called microcephaly, which prevents the brain from developing properly and results in individuals having a small head. Future work that builds on the findings presented by Sydor et al. could therefore help researchers to understand the causes of microcephaly in patients.
https://doi.org/10.7554/eLife.37846.002Introduction
The centrosome is a membrane-less organelle whose major role is to organize, orient, and regulate the site of microtubule formation. In somatic dividing cells, the centrosome is critical for ensuring faithful and timely chromosome segregation and establishment of the correct cell division axis, whereas in non-dividing and differentiated cells, it is critical for cellular polarization and cilia formation (Conduit et al., 2015; Vertii et al., 2016). Centrosomes are essential for normal human development and health (Nigg and Holland, 2018). Loss of function mutations in centrosomal proteins, including components of the centriolar cartwheel, elongation machinery, appendages, and pericentriolar material, are responsible for developmental defects such as primary recessive microcephaly (Barbelanne and Tsang, 2014), primordial dwarfism (Care4Rare Canada Consortium et al., 2015; Rauch et al., 2008; Zheng et al., 2016b), and ciliopathies (Reiter and Leroux, 2017). Defects in centrosome number and structure are a major hallmark of tumorigenesis (Gönczy, 2015; de Cárcer and Malumbres, 2014; Nigg and Holland, 2018). Recently, studies in mouse models indicated that centrosome over-duplication concomitant with mutations in p53 drives tumor formation in the epidermis (Serçin et al., 2016) and can drive tumor formation in certain other tissues, even in the absence of concurrent p53-/- mutations (Levine et al., 2017). Therefore, it is essential to characterize the critical set of proteins required for centrosome assembly to understand the molecular mechanism of disease and identify therapeutic targets (Nigg and Holland, 2018).
Due to its important role in cell and tissue homeostasis, the centrosome is built in a highly-regulated, stepwise manner through the assembly of a multiplicity of protein complexes (Conduit et al., 2015; Mennella et al., 2014). Significant progress has been made in understanding how centrosome duplication begins in most somatic cells—at the G1/S phase boundary—with the assembly of the cartwheel, a nine-fold symmetrical scaffold made of SAS6, STIL, and CEP135. While SAS6 molecules can undergo remarkable self-assembly in vitro, the kinase Plk4 promotes cartwheel formation and centriole duplication by phosphorylating STIL to favor its interaction with SAS6 (Vulprecht et al., 2012; Lin et al., 2013b; Dzhindzhev et al., 2014; Arquint and Nigg, 2016). The initial binding of Plk4 to the centriole is governed by CEP63 (Brown et al., 2013), CEP152 (Brown et al., 2013; Kim et al., 2013; Sonnen et al., 2013; Dzhindzhev et al., 2010; Hatch et al., 2010; Cizmecioglu et al., 2010), and CEP192 (Kim et al., 2013; Sonnen et al., 2013). After cartwheel formation, CPAP, recruited by STIL (Tang et al., 2011), aids in the formation of the centriole microtubule wall (Pelletier et al., 2006; Schmidt et al., 2009) by regulating centriolar microtubule plus-end dynamics (Basten and Giles, 2013; Zheng et al., 2016a). CEP135 facilitates the stabilization of the centriole structure (Ohta et al., 2002; Basten and Giles, 2013) but may also play a more direct role in initial cartwheel formation as recombinant Drosophila SAS6 and Bld10 (Drosophila CEP135 homolog) can self-organize into a nine-fold symmetrical cartwheel structure (Guichard et al., 2017).
Once the initial steps of procentriole formation occur, centriole elongation can proceed. However, we have a limited understanding of the essential components required for centriole elongation, which happens between S and G2 phases, and how they are assembled in a stepwise manner. CPAP has been shown to interact with CEP120 (Lin et al., 2013a) and SPICE (Comartin et al., 2013) in a complex that regulates centriole elongation at the centriolar microtubule wall (Archinti et al., 2010; Lin et al., 2013b). Centrobin has also been implicated in directly regulating centriolar microtubule elongation (Lee et al., 2010; Zou et al., 2005) and stability by binding to α-Tubulin (Gudi et al., 2011) and by regulating CPAP levels (Gudi et al., 2015; Gudi et al., 2014). Centrobin is further required to recruit CP110, a protein forming a cap-like structure on the distal end of the centriole that suppresses centriole elongation (Schmidt et al., 2009). Proximal to CP110, several proteins localized to the distal luminal end of centrioles such as POC5 (Azimzadeh et al., 2009), POC1B (Venoux et al., 2013), and OFD1 (Singla et al., 2010) have been implicated in promoting the elongation of the centriole’s distal region. More recently, additional proteins have been identified, namely CEP295 (Chang et al., 2016) and RTTN (Chen et al., 2017), which have been proposed to play a scaffolding role in the elongation process by connecting the centriole wall to the luminal centriolar region. However, it remains unclear if there are components in the lumen of the centriole that stabilize interactions with centriolar wall proteins.
RTTN (rotatin) was originally identified as a protein critical for axial rotation and left-right symmetry specification in mice (Faisst et al., 2002). Subsequently, mutations in human RTTN have been shown to cause primary microcephaly and primordial dwarfism (Kheradmand Kia et al., 2012; Grandone et al., 2016; Care4Rare Canada Consortium et al., 2015). Recent reports have shed light on the cellular function of RTTN. The Drosophila RTTN homolog, Ana3, was demonstrated to be a centrosomal component critical for maintaining the structural integrity of centrioles (Stevens et al., 2009), whereas human RTTN, localized near the centriolar cartwheel, has been shown to be dispensable for initial centriole assembly, but critical for formation of a full-length centriole (Chen et al., 2017). It remains unclear what factors are downstream of RTTN and how they promote the elongation and stabilization of the centriole once the cartwheel is formed.
Here, we characterize human PPP1R35, the product of the gene C7orf47, which was previously identified in fractions co-purifying with centrosomes in a high-throughput mass spectrometry study (Jakobsen et al., 2011). Our study demonstrates that PPP1R35 is a centrosomal component located in the proximal centriolar lumen above the cartwheel. We further demonstrate that PPP1R35 is not important for early centriole assembly but is critical for centriole elongation by impacting the recruitment of the microtubule-binding elongation machinery. In addition, we show that PPP1R35 is downstream of RTTN in the elongation pathway and that they form a protein complex. Altogether, we describe a novel centriolar component essential for centriole formation and identify a new mechanistic step downstream of RTTN in the pathway to reach a fully elongated centriole and functional centrosome.
Results
PPP1R35 is stably associated with the centrosome
To examine if PPP1R35 is a bona fide centrosomal protein, we generated a U2OS cell line constitutively expressing GFP-PPP1R35 under the control of a low copy protein expression promoter (Kim et al., 2011), integrated into the genome through the Flp-In system. GFP-PPP1R35 showed two main protein populations: one enriched in a diffraction limited spot located in the middle of the cell adjacent to the nucleus, consistent with centrosomal localization, and a cytoplasmic pool (Figure 1). To verify that the observed PPP1R35 was located at the centrosomes, we transfected the GFP-PPP1R35 U2OS Flp-In cell line with a vector expressing Centrin 1-mCherry and observed co-localization of Centrin 1-mCherry with GFP-PPP1R35 (Figure 1a). To examine the dynamics of PPP1R35 during the cell cycle, we conducted long-term live-cell imaging by spinning disc confocal fluorescence microscopy (Figure 1b and Video 1). PPP1R35 was found on two centrosomes (grandmother and mother) throughout the entire cell cycle (Figure 1b). We observed some cells that have four GFP-PPP1R35 spots prior to mitosis (Video 1) and noted that a second GFP-PPP1R35 spot was always present after mitosis, suggesting that PPP1R35 is recruited on daughter centrioles prior to mitosis. To verify that both the mother and daughter centrioles have recruited GFP-PPP1R35, we leveraged the ~1.5 x resolution increase of sub-diffraction live cell imaging. In all G2 cells examined, prior to centrosome separation, two GFP-PPP1R35 spots are resolvable on each of the centrioles (grandmother and mother), confirming that PPP1R35 resides on both the mother and daughter centrioles and must be recruited early in the duplication cycle, in S or early G2 phase (Figure 1c). To confirm that GFP-PPP1R35 localization is consistent with the endogenous protein, we imaged U2OS cells labeled with antibodies against PPP1R35 and γ-tubulin by confocal microscopy (Figure 1d) and cells labeled with antibodies against PPP1R35 and CETN1 by 3D structured illumination microscopy (3DSIM), and observed co-localization (Figure 1e). Since the anti-PPP1R35 antibody showed high background staining we used GFP-PPP1R35 to conduct further studies. To ensure that the GFP tag did not impact the localization of the protein, we examined the morphology of the centrosome by 3DSIM and did not observe a difference between WT and GFP-PPP1R35-expressing U2OS cells (Figure 1—figure supplement 1a). Furthermore, we verified that the GFP-PPP1R35 construct did not alter centrosome biogenesis by measuring the total number of CEP152-labeled centrosomes in WT and GFP-PPP1R35-expressing U2OS cells (Figure 1—figure supplement 1b). In addition, the GFP-PPP1R35 construct rescues the centriole duplication phenotype when PPP1R35 levels are knocked down by siRNA targeting the 3’ untranslated region (3’UTR, see below Figure 3c).
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Figure 1—source data 1
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Next, to determine whether PPP1R35 was continuously recruited or was stably associated to the centrosome, we performed Fluorescence Recovery After Photobleaching (FRAP) experiments. Comparison of the fluorescence recovery curves of the cytoplasmic versus centrosomal PPP1R35 pools revealed that centrosomal PPP1R35 did not fully recover to pre-bleach levels after photobleaching, therefore indicating that the protein has low turnover and is stably associated at the centrosome (Figure 1f,g). This observation is consistent with a previous analysis that identified PPP1R35 as co-purifying with centrosomal components and observed only a 22% turnover in centrosomal PPP1R35 as measured by stable isotope labeling of amino acids in cell culture (SILAC) mass spectrometry (Jakobsen et al., 2011). Altogether, our imaging experiments demonstrate that PPP1R35 is a resident protein of the centrosome and is recruited to the nascent daughter centriole early in the duplication cycle.
PPP1R35 localizes to the under-characterized proximal centriolar lumen above the cartwheel
To further dissect the role of PPP1R35 at the centrosome, we used super-resolution microscopy to precisely map the position of PPP1R35 relative to several reference markers, whose position at the centrosome has been previously characterized by EM and fluorescence imaging (Figure 2). To perform these experiments, we used linear 3DSIM, a technique that provides a 2-fold resolution increase over standard confocal/widefield fluorescence microscopy, which is sufficient to resolve the relative distribution of many centrosomal proteins and allows for straightforward multicolor imaging (Sydor et al., 2015; Mennella et al., 2012). 3DSIM imaging showed that GFP-PPP1R35 is located in the centrosomal lumen, as suggested by the position of PPP1R35 in the middle of the ring structure formed by CEP152 (Hatch et al., 2010; Cizmecioglu et al., 2010) (Figure 2a). Next, we used several proximal (SAS6, CEP135, CPAP, CEP250) and distal (CETN1, POC1B, POC5) proteins to locate the position of PPP1R35 along the centrosomal longitudinal axis. Qualitative assessment of the 3DSIM micrographs showed that the position of PPP1R35 is biased toward proximal markers such as CPAP and CEP135 more than either the utmost proximal (i.e. CEP250) or distal (i.e. CETN1) ends of the centriole (Figure 2b). To precisely map PPP1R35, we performed a quantitative analysis of the distance between PPP1R35 and many centriolar reference markers. We collected hundreds of 3DSIM images and analyzed micrographs with centriole side views where PPP1R35 was in the same z-plane of the protein of interest for measurement to avoid distortions due to anisotropic resolution (Figure 2c). 3DSIM molecular mapping shows that PPP1R35 is located furthest from the distal end proteins (Centrin-1: 230 ± 50 nm; POC5: 160 ± 50 nm; POC1B: 140 ± 60 nm), but closer to proximal end markers such as CEP135 (90 ± 40 nm) and CPAP (60 ± 30 nm), yet not as proximal as SAS6 (120 ± 40 nm) or CEP250 (170 ± 50 nm) (Figure 2d). Together, we conclude that PPP1R35 localizes to the proximal centriolar lumen just above the cartwheel (Figure 2e).
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Figure 2—source data 1
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PPP1R35 is critical for centriole component recruitment
Since PPP1R35 is recruited early in the centrosome duplication pathway, we hypothesized that it might play a role in regulating centrosome biogenesis. To test this possibility, we depleted PPP1R35 protein levels in U2OS cells by targeting the mRNA with two non-overlapping siRNA strands, one designed to be complementary to an exon in the conserved C-terminal region and the second to the 3’ UTR of the PPP1R35 mRNA (Figure 3a,b). The specificity of the siRNA strands toward PPP1R35 was validated by western blotting of cells expressing GFP-PPP1R35 (Figure 3—figure supplement 1) and RT-qPCR (Figure 3—figure supplement 2). We opted to deplete cells of PPP1R35 via siRNA rather than CRISPR/Cas9 gene editing since previous studies demonstrated cell lethality in the absence of PPP1R35 (Hart et al., 2015; Neumann et al., 2010). Cells were treated with siRNA for 72 hr, thereby allowing cells to progress through multiple cell cycles and accumulate any centriolar defects (Figure 3b and Figure 3—figure supplement 3). With both siRNA strands, a significant decrease was observed in centrosomal staining of CEP152, a protein recruited in the last stages of daughter centriole formation (Fu et al., 2016)(Figure 3c). This phenotype is rescued by exogenously expressing GFP-PPP1R35, demonstrating the specificity of the siRNA and the resultant phenotype of PPP1R35 loss (Figure 3c). We next sought to narrow down the stage of centrosome duplication at which PPP1R35 plays a role by labeling PPP1R35-depleted cells with several centrosomal proteins sequentially recruited during its assembly (Conduit et al., 2015; Fu et al., 2015; Loncarek and Bettencourt-Dias, 2018). This analysis revealed that centriolar components recruited early in the pathway such as SAS6 (Dzhindzhev et al., 2014), CEP135 (Loncarek and Bettencourt-Dias, 2018; Fu et al., 2015) and Centrin 1 (Middendorp et al., 1997), are modestly affected at centrioles in the absence of PPP1R35, as opposed to proteins recruited in later stages, such as CEP295 (Chang et al., 2016), POC1B (Venoux et al., 2013), and CEP152 (Loncarek and Bettencourt-Dias, 2018; Fu et al., 2015) that are drastically reduced (Figure 3e and Figure 3—figure supplement 4). To assay for centrosome function, we examined whether centrosomes could recruit the pericentriolar material or efficiently nucleate microtubules in the absence of PPP1R35 by staining for Cdk5rap2 and γ-tubulin. In both cases, we observed a significant reduction upon PPP1R35 knockdown (Figure 3—figure supplement 5).
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Figure 3—source data 1
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A more significant impact on centriolar components recruited later in the pathway is more noticeable in a time-course experiment, in which cells are assayed at 24 hr intervals after siRNA treatment (Figure 3f and Figure 3—figure supplement 3). In these assays, there is little change in the recruitment of early-centriolar components such as SAS6, CETN1, and CEP135 up to the 72 hr time point. In contrast, defective recruitment of other components, such as CPAP and CEP152, is present around the 48 hr time point. When cells treated with PPP1R35 siRNA were stained for CEP152 and SAS6, the proportion of engaged centrosomes with cartwheels was not significantly different (Figure 3g), further suggesting that PPP1R35 loss does not influence the early stages of centriole biogenesis. It is also noteworthy that at longer timepoints (>72hr), CETN1 levels drastically decrease suggesting that overall centriole formation is being impacted. Altogether, these results demonstrate that PPP1R35 loss of function results in decreased centrosome number and suggest that PPP1R35 is critical for the recruitment of centriolar components after cartwheel formation.
Biotinylation-dependent proximity mapping of PPP1R35 identifies the microcephaly protein RTTN
To better understand the mechanistic role of PPP1R35 in centriole duplication, we conducted biotinylation-dependent proximity mapping (BioID) (Roux et al., 2012) experiments using stable cell lines expressing protein fusions with a FLAG-BirA (R118G) (henceforth referred to as BirA*) tag on either the N- or C-terminus of PPP1R35. BioID analysis revealed a proximity map with several high-confidence hits (FDR score <1%). As expected, the proximity interactome of PPP1R35 shows several centrosomal proteins, including AZI1 (CEP131), CEP85, and KIAA0753 (Moonraker). One of the most robust hits, as evidenced by the high numbers of peptides identified by both N- and C-terminal BirA* tags, is RTTN, a recently characterized protein (H.-Y. Chen et al., 2017) whose mutations in patients cause microcephaly (Care4Rare Canada Consortium et al., 2015; Grandone et al., 2016), dwarfism, and polymicrogyria (Kheradmand Kia et al., 2012) (Figure 4a; Supplementary file 1). Stable HEK293T T-REX Flp-In cell lines showed normal centriolar localization as determined via confocal imaging with the marker CEP152 (Figure 5f).
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Figure 4—source data 1
- https://doi.org/10.7554/eLife.37846.023
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Figure 5—source data 1
- https://doi.org/10.7554/eLife.37846.030
PPP1R35 forms a complex with RTTN and the two proteins are mutually required for each other’s recruitment
We reasoned that if PPP1R35 and RTTN are in close proximity to each other and are both located at the centrosome near the cartwheel (H.-Y. Chen et al., 2017), they might form a bona fide protein complex. To test this hypothesis, we performed FLAG immunoprecipitation (IP) of the N- and C-terminal tagged PPP1R35 constructs (Figure 4b and Supplementary file 1). Notably, RTTN was found to form a complex with the N-terminally tagged BirA*-PPP1R35. RTTN was the only protein identified with high confidence by BioID that also co-immunoprecipitated with PPP1R35, suggesting a strong link between this microcephaly protein and PPP1R35 function. Interestingly, IP-mass spectrometry data using the N-terminal FLAG-tagged PPP1R35, but not the C-terminal FLAG-tag construct, detected a high-confidence interaction with RTTN. IP with the PPP1R35 C-terminal FLAG-tag construct still identified RTTN peptide counts above that of the controls, but below our confidence level cut-off (Supplementary file 1), indicating that the interaction between the two proteins has been severely impaired but not completely abolished. Altogether, this suggests that the binding site might reside within the conserved C-terminal region of PPP1R35.
RTTN is a 298 kDa protein predicted to have an elongated, solenoid conformation (Fournier et al., 2013) that has been recently reported to localize to basal bodies (Stevens et al., 2009; Kheradmand Kia et al., 2012) and the centrosome (H.-Y. Chen et al., 2017; Stevens et al., 2009; Care4Rare Canada Consortium et al., 2015). Specifically, RTTN has been shown to localize to the proximal lumen of centrioles near CEP135 and the cartwheel (H.-Y. Chen et al., 2017). To further characterize the structural and functional relationship of PPP1R35 and the microcephaly protein RTTN, we mapped the position of RTTN relative to PPP1R35 by 3DSIM imaging and quantitative analysis (Figure 4c). To detect RTTN we used both an N-terminal mCherry-RTTN construct and an antibody recognizing residues 50–150 of RTTN. Our data show that RTTN localizes to the proximal centriole and it is located in close proximity to PPP1R35, consistent with our BioID findings (PPP1R35 distance from mCherry-RTTN, 80 ± 50 nm; anti-RTTN, 110 ± 50 nm; Figure 4d).
To further explore the functional relationship between PPP1R35 and RTTN, we depleted PPP1R35 from U2OS cells by siRNA and examined RTTN recruitment. The presence of RTTN at the centrosome is moderately, yet significantly, diminished upon PPP1R35 depletion (Figure 4e). When the reciprocal recruitment was explored by RTTN depletion, a major reduction in centrosomal PPP1R35 was observed (Figure 4f). This phenotype appears to be unrelated to the decrease in centriole number expected as a result of RTTN knockdown (Chen et al., 2017), because the number of cells with at least 2 centrin spots remains unchanged (Figure 4—figure supplement 1), yet the number of cells lacking GFP-PPP1R35 is significantly reduced. These results show that the two proteins co-localize at the centriole and that both proteins are mutually required for each other’s recruitment to the centriole, with RTTN exerting a more significant impact on PPP1R35 recruitment to the centriole. Altogether, our data suggest that RTTN and PPP1R35 form a complex and that RTTN acts upstream of PPP1R35.
Conserved serine phosphorylation sites and the canonical PP1-binding site in PPP1R35 are not critical for centrosome duplication
PPP1R35 is a highly conserved protein whose homologs are found across a wide range of eukaryotic species, ranging from the simple multicellular organism Trichoplax adhaerens to Homo sapiens (Figure 5—figure supplement 1). Interestingly, PPP1R35 homologues are found only in Holozoa species, correlating well with species presenting centrosomes, with the exception of Caenorhabditis elegans (Hodges et al., 2010). PPP1R35 can be divided into two major domains based on amino acid sequence homology: the highly divergent N-terminal domain and the more conserved C-terminal domain (Figure 5—figure supplement 2). Despite its variability across evolution, the N-terminus contains several highly conserved residues in mammalian species including three serine residues (S45, S47, S52 in Homo sapiens PPP1R35) previously found phosphorylated in large scale phospho-proteomic studies in both human and mouse cells (Olsen et al., 2010; Dephoure et al., 2008; Chi et al., 2008) (Figure 5a and Figure 5—figure supplement 2). In particular, S47 and S52 have been reported to be Cdk phosphorylation sites (Chi et al., 2008) (Figure 5a). As such, we hypothesized that these residues could be candidates for regulating PPP1R35 activity during centrosome duplication, as Cdk2 ensures that centrosome duplication takes place concomitantly with DNA synthesis in S-phase (Fu et al., 2015).
To probe the importance of these residues in the interaction with RTTN, we mutated all three serine residues to either non-phosphorylatable alanines (S45A, S47A, S52A) or to phospho-mimetic aspartic acids (S45D, S47D, S52D) and generated inducible HEK293 T-Rex Flp-In cell lines expressing the mutant N-terminal BirA*-PPP1R35 constructs. Neither the triple alanine nor the triple aspartic acid mutant significantly impacted the presence of PPP1R35 at the centrosome, nor its proximity to RTTN (Figure 5b,g). Furthermore, co-IP demonstrated that neither phospho-mutant impacted the interaction between PPP1R35 and RTTN (Figure 5—figure supplement 3).
We further evaluated the role of PPP1R35 phosphorylation on centriole duplication by examining whether the above phospho-mutants are able to rescue our centriole defect phenotype. When cells were depleted of endogenous PPP1R35 by the 3’ UTR-targeting siRNA and expressed the triple aspartic acid mutant (S45D, S47D, S52D) GFP-PPP1R35 in trans, we did not observe any reduction in centrosome number (Figure 5c). Despite multiple attempts, we were unable to generate a triple alanine (S45A, S47A, S52A) mutant cell line in U2OS cells, therefore we examined both triple mutant cell lines in HEK293 cells (Figure 5—figure supplement 4). Analysis of individual alanine mutants (S45A and S47A) in U2OS cells is also consistent with the notion that that despite their conservation, these resides are not playing a critical for PPP1R35’s function in centriole biogenesis (Figure 5c).
PPP1R35 is predicted to contain a canonical RVxF PP1-binding site (Peti et al., 2013), encompassing residues 77–81 (Hendrickx et al., 2009) in the N-terminal domain. This site is conserved only among Chordata species (Figure 5—figure supplement 1 and Figure 5—figure supplement 2). Interestingly, disruption of the predicted PP1 binding site by mutating two conserved residues, V79 and F81, to alanine (Peti et al., 2013) leads to proper targeting to the centriole (Figure 5g) and does not disrupt PPP1R35’s proximity to, or interaction with, RTTN (Figure 5d, Figure 5—figure supplement 3). Furthermore, the V79A, F81A mutant nearly completely rescued our centriole duplication phenotype (Figure 5e), suggesting that it is not critical for centriole biogenesis.
PPP1R35 loss results in shortened centrioles by preventing the recruitment of proteins responsible for centriole elongation
Since PPP1R35 forms a complex with the microcephaly protein RTTN and this protein has been previously linked to centriole elongation, where its loss resulted in shortened centrioles (Chen et al., 2017), we investigated whether PPP1R35 knockdown results in diminished centriole length. To this effect, we used 3DSIM to measure the distance between the proximal end of centrioles labeled with acetylated tubulin to CP110, which localizes to the centriole’s distal end (Figure 6a). Acetylated tubulin has been suggested to be an early tubulin modification during centriole duplication (Balashova et al., 2009) and it has been previously used for conducting centriole length measurements (Chen et al., 2017). Furthermore, we verified that tubulin acetylation is unaffected when PPP1R35 is knocked down by siRNA (Figure 6—figure supplement 1). To ensure that we were examining mature centrioles, we focused our analysis only on mother centrioles in G2 phase cells in which a clear mother and daughter centriole were present. The length of mother centrioles was determined to be 356 ± 65 nm in control-RNAi treated cells, in agreement with previous reports (Thauvin-Robinet et al., 2014). When PPP1R35 levels are knocked down by siRNA, we see a significant reduction in centriole length to 263 ± 69 and 246 ± 83 nm for the exon and 3’ UTR siRNA, respectively (Figure 6b). On the contrary, overexpression of PPP1R35 does not significantly change centriole length (382 ± 89 nm). Due to the small effect observed on RTTN recruitment when PPP1R35 levels are reduced, we hypothesized that the shorter centriole length may be due to the inability of nascent centrioles to recruit proteins involved in elongation. We then examined cells treated with PPP1R35 siRNA for recruitment of proteins involved in either microtubule stabilization/recruitment such as CPAP and SPICE (Archinti et al., 2010; Zheng et al., 2016a; Tang et al., 2009; Comartin et al., 2013; Lin et al., 2013b) or the elongation of the distal portion of the centriole such as POC5 (Azimzadeh et al., 2009) and both proteins were significantly reduced in the absence of PPP1R35. Consistently, CP110, a negative regulator of centriole elongation recruited early in the elongation pathway, was not significantly changed relative to control RNAi-treated cells (Figure 6c). Altogether, this demonstrates that PPP1R35 is a critical factor for centriole assembly by promoting recruitment of centriole elongation proteins.
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Figure 6—source data 1
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Discussion
PPP1R35 was initially suggested to be a centrosomal protein by mass spectrometry studies that identified PPP1R35 as co-purifying with isolated centrosomes (Jakobsen et al., 2011). Here we show that the uncharacterized protein PPP1R35 is stably associated at the centrosome throughout the cell cycle where it plays a critical role in its elongation. Our loss of function analysis places PPP1R35 relatively early in the centriole duplication pathway, after cartwheel formation and before complete centriole elongation. In addition, we demonstrate that the ultimate downstream effect of PPP1R35 loss is shortened centrioles, suggesting that PPP1R35 directly controls the elongation pathway. It is interesting to note that this diminished centriole duplication defect takes several days to become very pronounced suggesting that either the short centrioles are still competent to duplicate but to a diminished degree, or that only low levels of PPP1R35 are needed for proper biological activity. Phenotypic analysis places PPP1R35 upstream of CPAP, CEP295, SPICE and POC5, which are all proteins involved in centriole elongation (Tang et al., 2009; Lin et al., 2013b; Comartin et al., 2013; Chang et al., 2016), but downstream of RTTN, which also affects the recruitment of POC1B, POC5, and CEP295 similarly to PPP1R35 (Chen et al., 2017). Furthermore, BioID, IP, and 3DSIM data show that PPP1R35 and RTTN form a protein complex and that this complex is likely a result of a direct interaction as RTTN is the only protein identified in both BioID and IP experiments.
Whereas several distal luminal proteins have been reported to date (Azimzadeh et al., 2009; Venoux et al., 2013; Paoletti et al., 1996), PPP1R35 is the first to be mapped to the proximal luminal region, an uncharacterized ‘outpost’ right above the cartwheel. Since most proteins involved in centriole elongation are localized along the microtubule wall or the distal end of the centriole (Comartin et al., 2013; Hatzopoulos et al., 2013; Azimzadeh et al., 2009; Schmidt et al., 2009), the interesting distribution of PPP1R35 suggests that it likely acts directly through RTTN, which our imaging show luminal localization for the N-terminus of the protein, in close proximity to PPP1R35.
Our phylogenetic analysis demonstrates that PPP1R35 is conserved in a wide range of species, including Drosophila, parasitic worms, and mammals (Figure 5—figure supplement 1). PPP1R35 is likely the human homolog of the Drosophila protein Reduction of Cnn dots 4 (Rcd4), a protein identified in a large RNAi screen aimed at discovering novel proteins impacting centrosome formation and PCM assembly (Dobbelaere et al., 2008). Alignment of Homo sapiens PPP1R35 and Drosophila Rcd4 results in an overall similarity of 24%, with the greatest homology in the conserved C-terminal domain (Figure 5—figure supplement 2). In all identified PPP1R35 homologs, the N-terminus exhibits a large degree of variability, hinting at an organism-specific specialization for this domain. We probed several conserved residues in PPP1R35 including several conserved serines (S45, S47, S52), but none of these mutants appeared to drastically impact centriole biogenesis. Intriguingly, the aforementioned serines and putative PP1-binding site, which are well conserved in Chordata, are not conserved in Rcd4. Furthermore, changes to the phosphorylation state did alter the overall BioID proximity map of PPP1R35, including altering the proximity to CEP85, AZI1, and OFD1, the latter two shown to have important roles in ciliogenesis (Hall et al., 2013; Ma and Jarman, 2011; Wilkinson et al., 2009; Romio et al., 2004; Ferrante et al., 2006).
The centrosomal duplication cycle is closely linked to the cell cycle and tightly controlled by a host of kinases and phosphatases (Pihan, 2013; Fujita et al., 2016). While kinases inherently possess temporal and spatial specificity, protein phosphatases require a regulatory component to properly function (Heroes et al., 2013; Peti et al., 2013; Korrodi-Gregório et al., 2014). To date, only a handful of centrosomal PP1 regulatory components have been identified (Katayama et al., 2001; Meraldi and Nigg, 2001; Mi et al., 2007; Helps et al., 2000; DeVaul et al., 2013; Huang et al., 2005) and overall knowledge of their interaction and role with PP1 is limited in scope. PPP1R35 is annotated to be a PP1-regulatory protein and contains a canonical PP1-binding site. Despite previous reports that demonstrated PP1-binding and inhibition (Hendrickx et al., 2009; Fardilha et al., 2011), we were unable to identify any PP1 isoform in our BioID or IP screens using HEK293 cycling cells. This is not completely surprising as previous studies have encountered similar difficulties in identifying interactions between protein phosphatases and their interactors, frequently due to the transient nature of binding (St-Denis et al., 2016). We tested the role of this interaction in regard to centriole duplication by mutating the canonical PP1-binding site but found that this interaction with PP1 does not appear to be critical for PPP1R35’s role at the centrosome. Overall, this suggests that despite PPP1R35’s annotation and previous demonstration as a PP1 regulator, this activity may not be related to centriole biogenesis. However, we cannot yet rule out the possibility that a second, non-canonical PP1-binding site is involved or that the PP1-regulating activity of PPP1R35 is required only in specific cellular functions not investigated here, such as ciliogenesis. Nonetheless, the large number of robust, non-centrosomal BioID hits suggests that PPP1R35 may serve other functions in the cell apart from centriole duplication and perhaps these other functions require PPP1R35’s PP1-regulation activity.
Despite the importance of centriole elongation to numerous human diseases, the exact mechanisms through which elongation takes place is still poorly understood (Loncarek and Bettencourt-Dias, 2018). To date, two major pathways governing centriole elongation have been described, one positive-growth mechanism acting on assisting microtubule elongation (CPAP/CEP120/SPICE) (Kohlmaier et al., 2009; Schmidt et al., 2009; Tang et al., 2009; Lin et al., 2013b; Comartin et al., 2013) and a second negative-growth mechanism involving CP110 and CEP97, which form a cap-like structure on the centriole to restrict microtubule growth (Schmidt et al., 2009; Spektor et al., 2007; Franz et al., 2013; Chen et al., 2002). However, even with the discovery of additional proteins such as POC5 (Azimzadeh et al., 2009), CEP295 (Chang et al., 2016), and Centrobin (Gudi et al., 2015), all of which impact centriole elongation, we apparently have yet to acquire a complete picture of this process. Here, we have identified a novel key player of this process, PPP1R35. Our data suggest that PPP1R35 primarily impacts the CPAP/CEP120/SPICE and RTTN/CEP295 pathways of centriole elongation (Figure 6d,e). Previously, RTTN was proposed to be critical for stabilizing the early procentriole containing STIL, CPAP, and SAS6 and for recruiting CEP295, which in turn can recruit POC5 and POC1B (Chen et al., 2017). Our data are consistent with a model where the impact on centriole elongation occurs primarily through PPP1R35’s interaction with RTTN. Such a role is consistent with the localization of both PPP1R35 and RTTN, which are uniquely positioned just above the cartwheel in the region where elongation following initial centriole formation would occur. PPP1R35 could be involved in either modulating RTTN’s turnover or interactions with other proteins. This reasoning is supported by the observation that RTTN loss has been shown to cause lack of proper CEP295, POC1B, and POC5 recruitment and an interdependency with CPAP and CEP135 (Chen et al., 2017), consistent with our observed phenotype when PPP1R35 is knocked down.
Mutations in numerous centriolar proteins have been linked to microcephaly (Barbelanne and Tsang, 2014; Kaindl et al., 2010), including components involved in centriole elongation such as CPAP (Leal et al., 2003) and RTTN (Care4Rare Canada Consortium et al., 2015; Grandone et al., 2016). The discovery of additional microcephaly proteins will aid in our understanding of the disease and assist in the development of future therapies. Our data show that that PPP1R35 impacts the process of centriole elongation through a close relationship with a known microcephaly protein, RTTN, therefore suggesting that PPP1R35 may be one such candidate microcephaly gene. Future DNA sequencing of microcephaly patients and animal model studies are needed to address this issue.
Materials and methods
Plasmids and molecular biology
Request a detailed protocolTables detailing primers (Supplementary file 2) and siRNA strands (Supplementary file 3) used in this study are available in the Supplemental Material section. The construct pIRES Centrin1 mCherry was a gift from Matthieu Piel (Addgene plasmid # 64338). For cloning experiments, PCR products were amplified from plasmid cDNA (PPP1R35 cDNA, MGC clone Image ID 4773899; RTTN cDNA, GE Dharmacon ORFeome cDNA 25914), verified for specificity of amplification on an agarose gel and purified using the PureLink PCR Purification kit (Thermo Fisher Scientific) or gel-extracted (Qiagen DNA Gel Extraction Kit) when necessary. All cloning experiments were conducted using Gibson Assembly (New England Biolabs) according to the manufacturer’s instructions. Site-directed mutagenesis for the single serine mutants was conducted using QuikChangeII XL Lightning site-directed mutagenesis kit (Agilent). Gene synthesis (Invitrogen GeneArt) was used to generate mutant PPP1R35 containing the triple Ala, Asp, and V79A, F81A mutations and were subsequently cloned into the GFP-containing FRT vector via Gibson Assembly as described above. Newly generated plasmid constructs were verified using Sanger Sequencing (ACGT Corp., Toronto).
Cell culture
Request a detailed protocolU2OS and U2OS Flp-In cells were cultured in McCoy’s 5A medium (Gibco) supplemented with 10% fetal bovine serum (Wisent) and 1 X antibiotic/antimycotic (Gibco, 100 units/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml amphotericin B). HEK 293T TREX cells were maintained in Dulbecco’s Modified Eagle’s Medium (Gibco) supplemented with 10% tetracycline-free fetal bovine serum (Wisent) and 1 X antibiotic/antimycotic. All cells were cultured at 37°C with 5% CO2 and routinely passed. The cells were tested routinely for mycoplasma contamination using Invitrogen’s Mycoplasm Detection Kit.
For cellular transfection of DNA plasmids, JetPrime (Polyplus) was used according to the manufacturer’s instructions. Cells were subsequently selected in the appropriate antibiotic (puromycin for constructs in U2OS Flp-In cells and hygromycin for constructs in HEK293 Flp-In TREX cells) to generate cell lines with stably integrated transgenes. For siRNA transfections, Lipofectamine RNAiMax (Invitrogen) was used according to the manufacturer’s instructions. All siRNA strands were transfected at a final concentration of 40 nM and cells were assayed 72 hr post transfection. Scrambled siRNA and siRNA targeting GAPDH were used as negative and positive controls, respectively. GAPDH control knockdown efficacy was monitored by immunofluorescence and by western blot. The number of labeled centrosomes per cell (classified into categories as either 0, 1, 2 or >2 labeled centrosome; see Figure 3d for example of centrosome spots imaged in cells and Figure 3—figure supplement 4 for example of categories of centrosomal counts) were manually counted and all siRNA experiments were conducted in at least triplicate except for the HEK293 siRNA experiments that were conducted in at least duplicate.
Live-cell imaging
Request a detailed protocolFor live-cell imaging, cells were seeded on KOH-washed coverglass (Electron Microscope Sciences) to reduce background fluorescence and subsequently left overnight to adhere. The standard culture media was replaced with DMEM medium lacking phenol red (Gibco) supplemented with 10% fetal bovine serum and 1 X antibiotic/antimycotic. Cells were imaged on a Zeiss Axio Observer Spinning-disc microscope equipped with Yogokawa spinning disk head, Phototronics EM CCD camera, and a 63x objective (NA = 1.4). The samples were maintained at 37°C with 5% CO2 during imaging in an incubation chamber. Automated acquisition of a 30 µm z-stack with a 0.75 µm step size every 40–45 min was obtained using the Zeiss Zen Blue software. During acquisition the lowest possible minimal laser power was used to avoid phototoxicity, resulting in movies of an average length of 14 hr.
Fluorescence recovery after photobleaching (FRAP) experiments
Request a detailed protocolThe cells were imaged on a Leica SP8 Confocal DMI6000 microscope equipped with a HyD detector and a 63x (NA = 1.4) oil objective. The samples were maintained at 37°C with 5% CO2 during imaging in an incubation chamber. Samples were bleached with a white light laser for approximately 1.5 s and the subsequent recovery monitored for an additional 26.5 s. The resultant plots were analyzed and fit to a single exponential curve using the build-in FRAP analysis function in the Leica Analysis Suite X software.
Immunofluorescence
Request a detailed protocolA table detailing all antibodies used in this study, including concentrations and suppliers, is available in the key resources table. Cells were plated onto coverslips (Electron Microscope Sciences; previously cleaned with KOH) and left overnight to adhere. The cells were treated with 0.02% w/v digitonin in PBS for 5 min at RT to remove the cytoplasmic population of PPP1R35 followed by fixation with −20°C methanol for 20 min. The cells were blocked for 1 hr using 5% FBS in PBS supplemented with 0.5% Tween-20. The cells were incubated with primary and secondary antibodies for 40 min each at RT. To detect specific primary antibodies, Alexa 488-, Alexa 568-, or Alexa 647-conjugated IgGs were used as secondary antibodies at a dilution of 1:1000 (Invitrogen). Cell nuclei were stained with Hoescht 33342 (Thermo Fisher). Cells were mounted with 0.5% n-propyl gallate in 80% glycerol mounting media.
3D structured illumination microscopy
Request a detailed protocol3DSIM data were collected using an ELYRA PS.1 (Carl Zeiss Microscopy) with a Plan-Apochromat 63x or 100x/1.4 Oil immersion objective lens with an additional 1.6x optovar. An Andor iXon 885 EMCCD camera was used to acquire images with 101 nm/slice z-stack intervals over a 5–10 µm thickness. The fluorophores were excited with 405, 488, 555 and 647 nm wavelengths and band-pass 420–480, 495–550, 570–620, long-pass 655 and 750 nm filters were used to collect the emission wavelengths. Laser powers at the objective focal plane of 52.6 mW in the 2–12% range, exposure time between 50–250 ms and EMCCD camera gain values between 5 and 50 were used during image acquisition. For each image field, grid excitation patterns were collected for five phases and three rotation angles (−75o; −15o, +45o). The raw data were reconstructed using the SIM module of ZEN Black Software (version 8.1) with noise filter values between −6 and −3. Channel alignment was conducted using calibrated file generated from super-resolution Tetraspec beads (Carl Zeiss Microscopy). If appropriate, whole-volume images or maximum intensity projections were exported as tiff files to be further analyzed in ImageJ/Fiji (NIH).
Protein mapping and centriole length measurements by 3DSIM
Request a detailed protocolTo measure the position of PPP1R35 relative to various centriolar markers, only 3DSIM images in which both the fluorescence maxima of PPP1R35 and the corresponding reference protein were on the same z-slice were analyzed. The distance between the peak maxima for the two markers were determined using the caliper function built in to the Zeiss Zen Black software (see Figure 2c for an example). The centriole length measurements were conducted in an identical manner using CP110 as a distal end marker and the acetylated tubulin signal as a proximal end marker (see Figure 6a for an example).
Proximity-dependent biotinylation
Request a detailed protocolBioID was conducted as previously described (Firat-Karalar and Stearns, 2015; Gupta et al., 2015). To generate stable cell lines expressing recombinant BirA fusion proteins for BioID experiments, HEK293 Flp-In T-Rex cells were co-transfected with the pcDNA5/FRT/TO PPP1R35-FLAG-BirA* or pcDNA5/FRT/TO FLAG-BirA*-PPP1R35 plasmid and Flp Recombinase Expression plasmid pOG44 in a 1:20 ratio, and then selected for multiple passages with increasing antibiotics concentrations to reach final concentrations of 400 µg/ml Hygromycin B (Invitrogen) and 15 µg/ml Blasticidin (Gibco, Thermo Fisher Scientific). HEK293 TREX Flp-In cells expressing the appropriate transgene were cultured until 90–100% confluency and treated for 24 hr with 1 µg/ml tetracycline to induce BirA expression and 50 µM biotin to allow biotinylation of proteins. HEK293T TREX Flp-In cells transfected with a vector containing either the N- or C-terminal FLAG-BirA* but no PPP1R35, were processed in parallel as controls.
Cells were collected, pelleted, and washed three times with PBS prior to freezing. Cell pellets were processed for Bio-ID and FLAG ImmunoPrecipitation (IP) experiments as described previously (Coyaud et al., 2015). Interactor classification: bona fide interactors were defined as high confidence protein identifications (ProteinProphet p>0.85) with a SAINT score ≥0.75, based on 4 independent MS runs. Histone hits were eliminated. Fold-change was calculated as described previously (Coyaud et al., 2015).
Western blot
Request a detailed protocolTotal cell lysates were collected using RIPA lysis buffer (Pierce) supplemented with mammalian protease inhibitor (BioBasic; 100 mM PMSF, 1 mM Bestatin, 1.5 mM Pepstatin A, 1.4 mM E-64, 0.08 mM Aprotinin, 1 mM Leupeptin) and cell debris pelleted by spinning for 30 min at 12,000 rpm. Protein concentrations were determined using a BCA protein assay kit (Pierce). Protein lysate containing ~15–30 µg of total protein was loaded onto well of 4–12% Bis-Tris gels (Invitrogen). Proteins were transferred to nitrocellulose membrane for 2 hr on an Invitrogen Bolt Minigel Apparatus at 10 V and blocked with 5% skim milk for 1 hr. Membranes were subsequently incubated with specific antibodies overnight at 4°C. Secondaries conjugated with HRP (Cell Signalling) were used at a 1:2000 dilution. Blots were developed using the ECL Chemiluminescent Substrate Kit (Invitrogen).
Real-time quantitative polymerase chain reaction (RT-qPCR)
Request a detailed protocolRNA was extracted from cells using the GeneJet RNA Purification kit (Thermo Scientific) and subsequently treated with the RapidOut DNA Removal kit (Thermo Scientific). Purified RNA was quantitated and only RNA with an A260/A280 ratio greater than 1.8 was used for reverse transcription with the BioRad iScript cDNA Synthesis kit with 1 µg of RNA as the template. All quantitative PCR was performed using a CFX Connect Real-Time System (BioRad) with SsoAdvanced Universal SYBR Green Supermix (BioRad) and 500 nM combined primer concentration per well. The relative expression of the target genes were normalized to RNA polymerase II and TATA binding protein transcript levels for each condition and then relative to expression in the scrambled siRNA-treated sample. Primer sequences can be found in Supplementary file 2. No-template and no-reverse transcriptase controls were run for each primer pair to confirm the lack of primer–dimer formation/DNA contamination and genomic DNA contamination, respectively. At least three biological replicates were run per condition. Data were analyzed using the CFX Maestro software (BioRad). All kits were conducted as per the manufacturer’s protocol.
Statistical analysis
Request a detailed protocolAll siRNA experiments were analyzed as 2 × 2 contingency tables in which all cells for a given population (i.e. cells with >1 CEP152 spots) were pooled for all replicates. To determine the p-values compared to the scrambled siRNA control for each dataset, Barnard’s Test was used in R with unpooled variances (package by Kamil Erguler; available at https://github.com/kerguler/Barnard) (Erguler, 2015). A summary of all statistics for the siRNA experiments can be found in Supplementary file 4. For all other statistical tests, the Student’s T-Test was used. Error bars represent the standard deviation for all replicates. For all figures, the following conventions were used: ns (p>0.05), * (p≤0.05), ** (p≤0.01), *** (p≤0.001), **** (p≤0.0001).
Phylogenetics
Request a detailed protocolThe NCBI protein database was queried with the search term ‘PPP1R35’ and all resultant hits were downloaded. For species with multiple annotated isoforms, the longest was selected. Any entries that were also annotated as a protein of known function (i.e. transposase, helicase, etc) were removed. Furthermore, only one organism per genus was selected to ensure broad coverage yet avoiding artifacts caused by over-sampled genera. All entries were from the Holozoa group of Eukaryotes. In order to ensure that no sequences from other major eukaryotic groups were missed, Delta Blastp searches using both the Homo sapien and Drosophila melanogaster PPP1R35 sequences were used to search for homologs in representative genera from the remaining eukaryotic groups (exact genera probed are those found in Figure 1 of Ref. [Hodges et al., 2010]). No additional homologs were identified outside of the Holozoa. Multiple sequence alignments were performed using Clustal Omega (Sievers et al., 2011) with the default settings. The phylogeny was inferred using the Bayesian method implemented with MrBayes v. 3.2.6 (mixed amino acid rate mode) and run for 2.5 million generations until the standard deviation of split frequencies was 0.199. Drosophila melanogaster Sds22, a PP1 regulator protein identified to have diverged early from homologous PP1 regulators (Ceulemans et al., 2002), was used as the outgroup. Trees were drawn using FigTree v. 1.4.3.
Data availability
All data generated or analyzed during this study are included in the manuscript and supporting files.
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Article and author information
Author details
Funding
National Science and Engineering Research Council of Canada (Discovery grant, RGPIN-2015-04795)
- Vito Mennella
The Hospital for Sick Children (Restracomp Postdoctoral Fellowship)
- Andrew Michael Sydor
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
Acknowledgements
We would like to thank the Trimble and Pelletier labs for vectors and reagents, Dr. Moshe Kim for the empty pcDNA5-FRT-TO- GFP Sept2p plasmid, the Mennella laboratory for insightful discussions and feedback, the National Science and Engineering Research Council of Canada for generous funding. Andrew Sydor is a Restracomp Fellow from the Hospital for Sick Children.
Copyright
© 2018, Sydor et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.
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Further reading
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- Cell Biology
Multiple gut antimicrobial mechanisms are coordinated in space and time to efficiently fight foodborne pathogens. In Drosophila melanogaster, production of reactive oxygen species (ROS) and antimicrobial peptides (AMPs) together with intestinal cell renewal play a key role in eliminating gut microbes. A complementary mechanism would be to isolate and treat pathogenic bacteria while allowing colonization by commensals. Using real-time imaging to follow the fate of ingested bacteria, we demonstrate that while commensal Lactiplantibacillus plantarum freely circulate within the intestinal lumen, pathogenic strains such as Erwinia carotovora or Bacillus thuringiensis, are blocked in the anterior midgut where they are rapidly eliminated by antimicrobial peptides. This sequestration of pathogenic bacteria in the anterior midgut requires the Duox enzyme in enterocytes, and both TrpA1 and Dh31 in enteroendocrine cells. Supplementing larval food with hCGRP, the human homolog of Dh31, is sufficient to block the bacteria, suggesting the existence of a conserved mechanism. While the immune deficiency (IMD) pathway is essential for eliminating the trapped bacteria, it is dispensable for the blockage. Genetic manipulations impairing bacterial compartmentalization result in abnormal colonization of posterior midgut regions by pathogenic bacteria. Despite a functional IMD pathway, this ectopic colonization leads to bacterial proliferation and larval death, demonstrating the critical role of bacteria anterior sequestration in larval defense. Our study reveals a temporal orchestration during which pathogenic bacteria, but not innocuous, are confined in the anterior part of the midgut in which they are eliminated in an IMD-pathway-dependent manner.
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- Cancer Biology
- Cell Biology
Understanding the cell cycle at the single-cell level is crucial for cellular biology and cancer research. While current methods using fluorescent markers have improved the study of adherent cells, non-adherent cells remain challenging. In this study, we addressed this gap by combining a specialized surface to enhance cell attachment, the FUCCI(CA)2 sensor, an automated image analysis pipeline, and a custom machine learning algorithm. This approach enabled precise measurement of cell cycle phase durations in non-adherent cells. This method was validated in acute myeloid leukemia cell lines NB4 and Kasumi-1, which have unique cell cycle characteristics, and we tested the impact of cell cycle-modulating drugs on NB4 cells. Our cell cycle analysis system, which is also compatible with adherent cells, is fully automated and freely available, providing detailed insights from hundreds of cells under various conditions. This report presents a valuable tool for advancing cancer research and drug development by enabling comprehensive, automated cell cycle analysis in both adherent and non-adherent cells.