Tag: multidisciplinary

  • Online images amplify gender bias

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    Here we outline the computational and experimental techniques we use to compare gender bias in online images and texts. We begin by describing the methods of data collection and analyses developed for the observational component of our study. Then we detail the study design deployed in our online search experiment. The preregistration for our online experiment is available at https://osf.io/3jhzx. Note that this study is a successful replication of a previous study with a nearly identical design, except the original study did not include a control condition nor several versions of the text condition; the preregistration of the previous study is available at https://osf.io/26kbr.

    Observational methods

    Data collection procedure for online images

    Our crowdsourcing methodology consisted of four steps (Extended Data Fig. 1). First, we gathered all social categories in WordNet, a canonical lexical database of English. WordNet contained 3,495 social categories, including occupations (such as ‘physicist’) and generic social roles (such as ‘colleague’). Second, we collected the images associated with each category from both Google and Wikipedia. Third, we used Python’s OpenCV—a popular open-source deep learning framework—to extract the faces from each image; this algorithm automatically isolates each face and extracts a square including the entire face and minimal surrounding context. Using OpenCV to extract faces helped us to ensure that each face in each image was separately classified in a standardized manner, and to avoid subjective biases in coders’ decisions for which face to focus on and categorize in each image. Fourth, we hired 6,392 human coders from MTurk to classify the gender of the faces. Following earlier work, each face was classified by three unique annotators16,17, so that the gender of each face (‘male’ or ‘female’) could be identified based on the majority (modal) gender classification across three coders (we also gave coders the option of labelling the gender of faces as ‘non-binary’, but this option was only chosen in 2% of cases, so we excluded these data from our main analyses and recollected all classifications until each face was associated with three unique coders using either the ‘male’ or the ‘female’ label). Although coders were asked to label the gender of the face presented, our measure is agnostic to which features the coders used to determine their gender classifications; they may have used facial features, as well as features relating to the aesthetics of expressed gender such as hair or accessories. Each search was implemented from a fresh Google account with no prior history. Searches were run in August 2020 by ten distinct data servers in New York City. This study was approved by the Institutional Review Board at the University of California, Berkeley, and all participants provided informed consent.

    To collect images from Google, we followed earlier work by retrieving the top 100 images that appeared when using each of the 3,495 categories to search for images using the public Google Images search engine16,17,18 (Google provides roughly 100 images for its initial search results). To collect images from Wikipedia, we identified the images associated with each social category in the 2021 Wikipedia-based Image Text Dataset (WIT)27. WIT maps all images across Wikipedia to textual descriptions on the basis of the title, content and metadata of the active Wikipedia articles in which they appear. WIT contained images associated with 1,523 social categories from WordNet across all English Wikipedia articles (see Supplementary Information section A.1.1 for details on our Wikipedia analysis). The coders identified 18% of images as not containing a human face; these were removed from our analyses. We also asked all annotators to complete an attention check, which involved choosing the correct answer to the common-sense question “What is the opposite of the word ‘down’?” from the following options: ‘Fish’, ‘Up’, ‘Monk’ and ‘Apple’. We removed the data from all annotators who failed an attention check (15%), and we continued collecting classifications until each image was associated with the judgements of three unique coders, all of whom passed the attention check.

    Collecting human judgements of social categories

    We hired a separate sample of 2,500 human coders from MTurk to complete a survey study in which they were presented with social categories (five categories per task) and asked to evaluate each category by means of the following question (each category was assessed by 20 unique human coders): “Which gender do you most expect to belong to this category?” This was answered as a scalar with a slider ranging from −1 (females) to 1 (males). All MTurkers were prescreened such that only US-based MTurkers who were fluent in English were invited to participate in this task.

    Demographics of human coders

    The human coders were all adults based in the USA who were fluent in English. Supplementary Table 1 indicates that our main results are robust to controlling for the demographic composition of our human coders. Among our coders, 44.2% identified as female, 50.6% as male and 3.2% as non-binary; the remainder preferred not to disclose. In terms of age, 42.6% identified as being 18–24 years, 22.9% as 25–34, 32.5% as 35–54, 1.6% as 55–74 and less than 1% as more than 75. In terms of race, 46.8% identified as Caucasian, 11.6% as African American, 17% as Asian, 9% as Hispanic and 10.3% as Native American; the remainder identified as either mixed race or preferred not to disclose. In terms of political ideology, 37.2% identified as conservative, 33.8% as liberal, 20.3% as independent and 3.9% as other; the remainder preferred not to disclose. In terms of annual income, 14.3% reported making less than US$10,000, 33.4% reported US$10,000–50,000, 22.7% reported US$50,000–75,000, 14.9% reported US$75,000–100,000, 10.5% reported US$100,000–150,000, 2.8% reported US$150,000–250,000 and less than 1% reported more than US$250,000; the remainder preferred not to disclose. In terms of the highest level of education acquired by each annotator, 2.7% selected ‘Below High School’, 17.5% selected ‘High School’, 29.2% selected ‘Technical/Community College’, 34.5% selected ‘Undergraduate degree’, 14.8% selected ‘Master’s degree’ and less than 1% selected ‘Doctorate degree’; the remainder preferred not to disclose.

    Constructing a gender dimension in word embedding space

    Our method for measuring gender associations in text relies on the fact that word embedding models use the frequency of co-occurrence among words in text (for example, whether they occur in the same sentence) to position words in an n-dimensional space, such that words that co-occur together more frequently are represented as closer together in this n-dimensional space. The ‘embedding’ for a given word refers to the specific position of this word in the n-dimensional space constructed by the model. The cosine distance between word embeddings in this vector space provides a robust measure of semantic similarity that is widely used to unpack the cultural meanings associated with categories13,22,31. To construct a gender dimension in word embedding space, we adopt the methodology recently developed by Kozlowski et al.22. In their paper, Kozlowski et al.22 construct a gender dimension in embedding space along which different categories can be positioned (for example, their analysis focuses on types of sport). They start by identifying two clustered regions in word embedding space corresponding to traditional representations of females and males, respectively. Specifically, the female cluster consists of the words ‘woman’, ‘her’, ‘she’, ‘female’ and ‘girl’, and the male cluster consists of the words ‘man’, ‘his’, ‘he’, ‘male’ and ‘boy’. Then, for each of the 3,495 social categories in WordNet, we calculated the average cosine distance between this category and both the female and the male clusters. Each category, therefore, was associated with two numbers: its cosine distance with the female cluster (averaged across its cosine distance with each term in the female cluster), and its cosine distance with the male cluster (averaged across its cosine distance with each term in the male cluster). Taking the difference between a category’s cosine distance with the female and male clusters allowed each category to be positioned along a −1 (female) to 1 (male) scale in embedding space. The category ‘aunt’, for instance, falls close to −1 along this scale, whereas the category ‘uncle’ falls close to 1 along this scale. Of the categories in WordNet, 2,986 of them were associated with embeddings in the 300-dimensional word2vec model of Google News, and could therefore be positioned along this scale. All of our results are robust to using different terms to construct the poles of this gender dimension (Supplementary Fig. 18). However, our main analyses use the same gender clusters as ref. 22.

    To compute distances between the vectors of social categories represented by bigrams (such as ‘professional dancer’), we used the Phrases class in the Gensim Python package, which provided a prebuilt function for identifying and calculating distances for bigram embeddings. This method works by identifying an n-dimensional vector of middle positions between the vectors corresponding separately to each word in the bigram (for example, ‘professional’ and ‘dancer’). This technique then treats this middle vector as the singular vector corresponding to the bigram ‘professional dancer’ and is thereby used to calculate distances from other category vectors. This same method was applied to the construction of embeddings for all bigram categories in all models.

    To maximize the similarity between our text-based and image-based measures of gender association, we adopted the following three techniques. First, we normalized our textual measure of gender associations using minimum–maximum normalization, which ensured that a compatible range of values was covered by both our text-based and image-based measures of gender association. This is helpful because the distribution of gender associations for the image-based measure stretched to both ends of the −1 to 1 continuum as a result of certain categories being associated with 100% female faces or 100% male faces. By contrast, although the textual measure described above contains a −1 (female) to 1 (male) scale, the most female category in our WordNet sample has a gender association of −0.42 (‘chairwoman’), and the most male category has a gender association of 0.33 (‘guy’). Normalization ensures that the distribution of gender associations in the image- and text-based measures both equally cover the −1 to 1 continuum, so that paired comparisons between these scales (matched at the category level) can directly examine the relative ranking of a category’s gender association in each measure. Minimum–maximum normalization is given by the following equation:

    $$\widetilde{{x}_{i}}=\frac{\left({x}_{i}-{x}_{\min }\right)}{\left({x}_{\max }-{x}_{\min }\right)}$$

    (1)

    where xi represents the gender association of category xi ([−1,1]), xmin represents the category with the lowest gender score, xmax represents the category with the highest gender score, and \(\widetilde{{x}_{i}}\) represents the normalized gender association of category xi. To preserve the −1 to 1 scale in applying minimum–maximum normalization, we applied this procedure separately for male-skewed categories (that is, all categories with a gender association above 0), such that xmin represents the least male of the male categories and xmax represents the most male of the male categories. We applied this same procedure to the female-skewed categories, except that, because the female scale is −1 to 0, xmin represents the most female of the female categories and xmax represents the least female. For this reason, after the 0–1 female scale was constructed, we multiplied the female scores by −1 so that −1 represented the most female of the female categories and 0 represented the least. We then appended the female-normalized (−1 to 0) and male-normalized (0 to 1) scales. Both the male and female scales before normalization contained categories with values within four decimal points of zero (|x| < 0.0001), such that this normalization technique had no effect of arbitrarily pushing certain categories towards 0. Instead, the above technique has the advantage of stretching out the text-based measure of gender association to ensure that a substantial fraction of categories reach all the way to the −1 female region and all the way to the 1 male region of the continuum, similar to the distribution of values for the image-based measure.

    Experimental methods

    Participant pool

    For this experiment, a nationally representative sample of participants (n = 600) was recruited from the popular crowdsourcing platform Prolific, which provides a vetted panel of high-quality human participants for online research. No statistical methods were used to determine this sample size. A total of 575 participants completed the task, exhibiting an attrition rate of 4.2%. We only examine data from participants who completed the experiment. Our main results report the outcomes associated with the Image, Text and Control conditions (n = 423); in the Supplementary Information, we report the results of an extra version of the Text condition involving the generic Google search bar (n = 150; Supplementary Fig. 26). We only examine data from participants who completed the task. To recruit a nationally representative sample, we used Prolific’s prescreening functionality designed to provide a nationally representative sample of the USA along the dimensions of sex, age and ethnicity. Participants were invited to partake in the study only if they were based in the USA, fluent English speakers and aged more than 18 years. A total of 50.8% of participants were female (no participants identified as non-binary). All participants provided informed consent before participating. This experiment was run on 5 March 2022.

    Participant experience

    Extended Data Fig. 2 presents a schematic of the full experimental design. This experiment was approved by the Institutional Review Board at the University of California, Berkeley. In this experiment, participants were randomized to one of four conditions: (1) the Image condition (in which they used the Google Image search engine to retrieve images of occupations), (2) the Google News Text condition (in which they used the Google News search engine, that is, news.google.com, to retrieve textual descriptions of occupations), (3) the Google Neutral Text condition (in which they used the generic Google search engine, that is, google.com, to retrieve textual descriptions of occupations) and (4) the Control condition (in which they were asked at random to use either Google Images or the neutral (standard) Google search engine to retrieve descriptions of random, non-gendered categories, such as ‘apple’). Note that, in the main text, we report the experimental results comparing the Image, Control and Google News Text conditions; we present the results concerning the Google Neutral Text condition as a robustness test in the Supplementary Information (Supplementary Fig. 26).

    After uploading a description for a given occupation, participants used a −1 (female) to 1 (male) scale to indicate which gender they most associate with this occupation. In this way, the scale participants used to indicate their gender associations was identical to the scale we used to measure gender associations in our observational analyses of online images and text. In the control condition, participants were asked to indicate which gender they associate with a given randomly selected occupation after uploading a description for an unrelated category. Participants in all conditions completed this sequence for 22 unique occupations (randomly sampled from a broader set of 54 occupations). These occupations were selected to include occupations from science, technology, engineering and mathematics, and the liberal arts. Each occupation that was used as a stimulus could also be associated with our observational data concerning the gender associations measured in images from Google Images and the texts of Google News. Here is the full preregistered list of occupations used as stimuli: immunologist, mathematician, harpist, painter, piano player, aeronautical engineer, applied scientist, geneticist, astrophysicist, professional dancer, fashion model, graphic designer, hygienist, educator, intelligence analyst, logician, intelligence agent, financial analyst, chief executive officer, clarinetist, chiropractor, computer expert, intellectual, climatologist, systems analyst, programmer, poet, astronaut, professor, automotive engineer, cardiologist, neurobiologist, English professor, number theorist, marine engineer, bookkeeper, dietician, model, trained nurse, cosmetic surgeon, fashion designer, nurse practitioner, art teacher, singer, interior decorator, media consultant, art student, dressmaker, English teacher, literary agent, social worker, screen actor, editor-in-chief, schoolteacher. The set of occupations that participants evaluated was identical across conditions.

    Once each participant completed this task for 22 occupations, they were then asked to complete an IAT designed to measure the implicit bias towards associating men with science and women with liberal arts33,34,35,38. The IAT was identical across conditions (‘Measuring implicit bias using the IAT’). In total, the experiment took participants approximately 35 minutes to complete. Participants were compensated at the rate of US $15 per hour for their participation.

    Measuring implicit bias using the IAT

    The IAT in our experiment was designed using the iatgen tool33 (https://iatgen.wordpress.com/). The IAT is a psychological research tool for measuring mental associations between target pairs (for example, different races or genders) and a category dimension (for example, positive–negative, science–liberal arts). Rather than measuring what people explicitly believe through self-report, the IAT measures what people mentally associate and how quickly they make these associations. The IAT has the following design (description borrowed from iatgen)33: “The IAT consists of seven ‘blocks’ (sets of trials). In each trial, participants see a stimulus word on the screen. Stimuli represent ‘targets’ (for example, insects and flowers) or the category (for example, pleasant–unpleasant). When stimuli appear, the participant ‘sorts’ the stimulus as rapidly as possible by pressing with either their left or right hand on the keyboard (in iatgen, the ‘E’ and ‘I’ keys). The sides with which one should press are indicated in the upper left and right corners of the screen. The response speed is measured in milliseconds.” For example, in some sections of our study, a participant might press with the left hand for all male + science stimuli and with their right hand for all female + liberal arts stimuli.

    The theory behind the IAT is that the participant will be fast at sorting in a manner that is consistent with one’s latent associations, which is expected to lead to greater cognitive fluency in one’s intuitive reactions. For example, the expectation is that someone will be faster when sorting flowers + pleasant stimuli with one hand and insects + unpleasant with the other, as this is (most likely) consistent with people’s implicit mental associations (example borrowed from iatgen). Yet, when the category pairings are flipped, people should have to engage in cognitive work to override their mental associations, and the task should be slower. The degree to which one is faster in one section or the other is a measure of one’s implicit bias.

    In our study, the target pairs we used were ‘male’ and ‘female’ (corresponding to gender), and the category dimension referred to science–liberal arts. To construct the IAT, we followed the design used by Rezaei38. For the male words in the pairs, we used the following terms: man, boy, father, male, grandpa, husband, son, uncle. For the female words in the pairs, we used the following terms: woman, girl, mother, female, grandma, wife, daughter, aunt. For the science category, we used the following words: biology, physics, chemistry, math, geology, astronomy, engineering, medicine, computing, artificial intelligence, statistics. For the liberal arts category, we used the following words: philosophy, humanities, arts, literature, English, music, history, poetry, fashion, film. Extended Data Figs. 3–6 illustrate the four main IAT blocks that participants completed (as per standard IAT design, participants were also shown blocks 2, 3 and 4, with the left–right arrangement of targets reversed). Participants completed seven blocks in total, sequentially. The IAT instructions for Extended Data Fig. 3 state, “Place your left and right index fingers on the E and I keys. At the top of the screen are 2 categories. In the task, words and/or images appear in the middle of the screen. When the word/image belongs to the category on the left, press the E key as fast as you can. When it belongs to the category on the right, press the I key as fast as you can. If you make an error, a red X will appear. Correct errors by hitting the other key. Please try to go as fast as you can while making as few errors as possible. When you are ready, please press the [Space] bar to begin.” These instructions are repeated throughout all blocks in the task.

    To measure implicit bias based on participants’ reaction times during the IAT, we adopted the following standard approach (used by iatgen). We combined the scores across all four blocks (blocks 3, 4, 6 and 7 in iatgen). Some participants are also faster than others, adding statistical ‘noise’ as a result of variance in overall reaction times. Thus, instead of comparing within-person differences in raw latencies, this difference is standardized at the participant level, dividing the within-person difference by a ‘pooled’ standard deviation. This pooled standard deviation uses the standard deviation of what are called the practice and critical blocks combined. This yields a D score. In iatgen, a positive D value indicates association in the form of target A + positive, target B + negative, which in our case is male + science, female + liberal arts), whereas a negative D value indicates the opposite bias (target A + negative, target B + positive, which in our case is male + liberal arts, female + science), and a zero score indicates no bias.

    Our main experimental results evaluate the relationship between the participants’ explicit and implicit gender associations and the strength of gender associations in the Google images and textual descriptions they encountered during the search task. The strength of participants’ explicit gender associations is calculated as the absolute value of the number they input using the −1 (female) to 1 (male) scale after each occupation they classified (Extended Data Fig. 2). Participants’ implicit bias is measured by the D score of their results on the IAT designed to detect associations between men and science and women and liberal arts. To measure the strength of gender associations in the Google images that participants encountered, we calculated the gender parity of the faces uploaded across all participants who classified a given occupation. For example, we identified the responses of all participants who provided image search results for the occupation ‘geneticist’, and we constructed the same gender dimensions as described in the main text, such that −1 represents 100% female faces, 0 represents 50% female (male) faces and 1 represents 100% male faces. To identify the gender of the faces of the images that participants uploaded, we recruited a separate panel of MTurk workers (n = 500) who classified each face (there were 3,300 images in total). Each face was classified by two unique MTurkers; if they disagreed in their gender assignment, a third MTurk worker was hired to provide a response, and the gender identified by the majority was selected. We adopted an analogous approach to annotating the gender of the textual descriptions that participants uploaded in the text condition. These annotators identified whether each textual or visual description uploaded by participants was female (1), neutral (0) or male (1). Each textual description was coded as male, female or neutral on the basis of whether it used male or female pronouns or names to describe the occupation (for example, referred to a ‘doctor’ as ‘he’); textual descriptions were identified as neutral if they did not ascribe a particular gender to the occupation described. We were then able to calculate the same measure of gender balance in the textual descriptions uploaded for each occupation as we applied in our image analysis.

    Reporting summary

    Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

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  • Room-temperature quantum optomechanics using an ultralow noise cavity

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    Fabrication of density-modulated membranes

    We use the soft clamping6,49,50,51,52,53 technique to realize ultrahigh mechanical quality factors. Our membrane design is inspired by those pioneered in ref. 7, but we use a different material for the nanopillars and a different fabrication process (see Supplementary Information for more details). We fabricated density-modulated PNC membranes by patterning amorphous silicon (aSi) nanopillars on a high aspect ratio Si3N4 membrane. In our PNC membranes, we fabricated pillars with diameters dpil = 300–800 nm, thickness of about 600 nm and nearest-neighbour distances apil = 1.0–2.0 μm. Amorphous silicon is grown with plasma-enhanced chemical vapour deposition (PECVD) at a temperature of 300 °C. Electron-beam lithography (FOx16 electron-beam resist) and dry etching (using a plasma of SF6 and C4F8) are used to pattern pillar arrays in aSi. Dry etching is stopped on a 6-nm layer of HfO2 (hafnium oxide) grown with atomic layer deposition (ALD) directly on top of Si3N4. HfO2 is used as an etch-stop layer because it is quite resistant to hydrofluoric acid (HF) etching, and the undercut created at the pillar base in the following process steps is limited. Undercut minimization is important to control the added dissipation induced by pillar motion (Supplementary Information). We remove the FOx mask and the residual etch-stop layer by dipping the wafer in HF 1% for about 3.5 min.

    After patterning the pillars, we encapsulate them in a PECVD SixNy layer to protect them during the silicon deep etching step. We first grow a thin (about 20 nm), protective layer of Al2O3 with ALD, to shield the membrane layer from plasma bombardment during PECVD. Then, approximately 125 nm of SixNy is grown at 300 °C, with 40 W of radio-frequency power exciting the plasma during deposition. This SixNy layer has been characterized to have a tensile stress of around +300 MPa at room temperature. The layer perfectly seals the nanopillars during immersion in hot KOH, without significant consumption.

    After patterning the pillars on the wafer frontside film, a thick (about 3 μm) layer of positive tone photoresist is spun on top for protection during the backside lithography process, which we perform with an MLA150 laser writer (Heidelberg Instruments). Optical lithography is followed by Si3N4 dry etching with a plasma of CHF3 and SF6. After the resist mask and protection layer removal with N-methyl-2-pyrrolidone (NMP) and O2 plasma, we deep-etch with KOH from the membrane windows while keeping the frontside protected, by installing the wafer in a watertight PEEK holder in which only the backside is exposed6. KOH 40% at 70 °C is used, and the etch is interrupted when about 30–40 μm of silicon remains. The wafer is then rinsed and cleaned with hot HCl of the residues formed during KOH etching. Then, the wafer is separated into individual dies with a dicing saw, and the process continues chipwise. Chips are again cleaned with NMP and O2 plasma, and the deep-etch is concluded with a second immersion in KOH 40% at a lower temperature of 55 °C, followed by cleaning in HCl. From the end of the KOH etching step, the composite membranes are suspended, and great care must be taken while displacing and immersing the samples in liquid. We dry the samples by moving them to an ultrapure isopropyl alcohol bath after water rinsing. Isopropyl alcohol has a high vapour pressure, and quickly evaporates from the chip interfaces, with few residues left behind.

    Finally, the PECVD nitride and Al2O3 layers can be removed selectively with wet etching in buffered HF. The chips are loaded in a Teflon carrier in which they are vertically mounted and immersed for about 3 min 20 s in BHF 7:1. It is crucial not to etch more than necessary to fully remove the encapsulation films: membranes become extremely fragile and the survival yield drops sharply when their thickness is reduced below around 15 nm. The membranes are then carefully rinsed, transferred in an ethanol bath and dried in a critical point dryer, in which the liquids can be evacuated gently and with little contamination.

    Fabrication and simulation of phononic-crystal-patterned mirrors

    The top and bottom mirror substrates are, respectively, fused silica and borosilicate glass, with a high-reflection coating sputtered on one face and an anti-reflection layer coating the other face. No layer for the protection of the optical coating is applied before machining. We use a dicing saw for glass machining to pattern a regular array of lines into the mirror substrates. The blade is continuously cooled by a pressurized water jet during the patterning process. The maximum cut depth allowed for our blade is 2.5 mm, and we constrain the designed PNC accordingly. We cut the flat bottom mirror from only one side (its thickness is only 1 mm), and the top mirror is patterned symmetrically with parallel cuts from both sides, as it is 4 mm thick. The relatively deep cuts in the top mirror need to be patterned over several passes, with gradually increasing depths. After patterning one mirror side, the piece is flipped and the other side is patterned after aligning to the first cuts, visible through the glass substrate. The lines are arranged in a square lattice for simplicity, although more complex patterns can be machined with the dicing saw. After the dicing process, the mirrors are subject to ultrasonic cleaning, while immersing first in acetone and then in isopropanol.

    We simulate the band diagrams of the unit cells of both the top and the bottom mirrors in COMSOL Multiphysics with the Structural Mechanics module. We optimized the lattice constant and cut depths to maximize the bandgap width, while centring the bandgap around 1 MHz and making sure that the remaining glass thickness is sufficient to maintain a reasonable level of structural stiffness. Details of the PNC dimensions are shown in the Supplementary Information. Owing to the finite size of the mirrors, we expect to observe edge modes within the mechanical bandgap frequency range. The thermal vibrations of these modes penetrate into the PNC structure with exponentially decaying amplitudes. To account for their noise contributions, we simulated the frequency noise spectrum of the MIM assembly (details shown in the Supplementary Information). The eigenfrequency solution confirmed the existence of edge modes with frequencies within the mechanical bandgap, but did not predict any significant contribution to the cavity frequency noise: the PNC is sufficiently large to reduce their amplitude at the cavity mode position.

    After patterning the PNC structures on the mirrors, we assembled a cavity with a spacer chip in place of a membrane and observed that the TE00 linewidth with the diced mirrors is identical to that of the original cavity. This indicates that our fabrication process does not cause measurable excess roughness or damage to the mirror surfaces. By contrast, when the assembly was clamped too tightly, excess cavity loss occurred because of significant deformation of the PNC mirrors, with a reduced stiffness. We mitigate this detrimental effect in the experiment by gently clamping the MIM cavity, with a spring compression sufficient to guarantee the structural stability of the assembly. We also ensure that the cavity mode is well-centred on the bottom mirror, to reduce the thermal noise contribution of the upper band-edge modes. For the MIM experiment discussed in the main text, we did not observe any mirror modes within the mechanical bandgap of the membrane chip. We can distinguish membrane modes from mirror modes by exploiting the fact that the coupling rates of membrane modes vary between different cavity resonances, whereas this is not the case for mirror modes.

    Nonlinear noise cancellation scheme

    At room temperature, the large thermal noise of the cavity, combined with the nonlinear cavity transduction response, results in a nonlinear mixing noise (TIN). This noise could lead to excess intracavity photon fluctuations and also to excess noise in optical detection. In the following, we discuss the strategy to cancel these effects in the fast-cavity limit (ωκ). Theoretical derivations and a discussion of the effect of a finite ω/κ ratio can be found in the Supplementary Information.

    In the experiment, we pump the cavity at the magic detuning, \(2\overline{\varDelta }/\kappa =-1/\sqrt{3}\), in which the nonlinear photon number noise is cancelled, to prevent excess oscillator heating due to nonlinear classical radiation pressure noise. To show the quantum correlations leading to optomechanical squeezing and conduct measurement-based state preparation, we need to perform measurements at arbitrary optical quadrature angles. Balanced homodyne detection provides the possibility of tuning the optical quadrature, but it does not offer enough degrees of freedom to cancel the nonlinear noise in detection. However, if the local oscillator is injected from a highly asymmetric beam splitter with a very small reflectivity (r 1) and the combined field is detected on a single photodiode, the photodetection nonlinearity is maintained and offers enough degrees of freedom to cancel the nonlinear noise in detection4 (for a derivation, see Supplementary Information). Specifically, simultaneous tuning of local oscillator amplitude and phase enables nonlinear mixing noise cancellation at arbitrary quadrature angles. In the fast-cavity limit, the cancellation condition is

    $$\begin{array}{l}\left|\frac{{\overline{a}}_{{\rm{sig}}}}{{\overline{a}}_{\hom }}\right|=2{\rm{Re}}\,\left[\frac{{{\rm{e}}}^{-{\rm{i}}\theta }}{{(-{\rm{i}}\overline{\varDelta }+\kappa /2)}^{2}}\right]\,\left[{\overline{\varDelta }}^{2}+{\left(\frac{\kappa }{2}\right)}^{2}\right]\\ \,=2\cos [\theta -2\arg ({\chi }_{{\rm{cav}}}(0))],\end{array}$$

    where \({\overline{a}}_{\hom }\approx {\overline{a}}_{{\rm{sig}}}+r{\overline{a}}_{{\rm{LO}}}\) is the coherent combination of the signal field \({\overline{a}}_{{\rm{sig}}}\) and the local oscillator field \({\overline{a}}_{{\rm{LO}}}\) (defined as the field before the beam splitter), θ = θhom − θsig is the quadrature rotation angle and \({\chi }_{{\rm{cav}}}(0)={\left(\kappa /2-i\overline{\varDelta }\right)}^{-1}\) is the cavity d.c. optical susceptibility.

    In the experiment, to detect a certain quadrature angle while cancelling nonlinear noise, we lock the homodyne power at the corresponding combined field level \({I}_{\hom }=| {\overline{a}}_{\hom }{| }^{2}\). We then continuously vary the local oscillator power using a tunable neutral density filter until the noise in the mechanical bandgap is perfectly cancelled. The level of mixing noise is very sensitive to the local oscillator power, and therefore the cancellation point can serve as a good indicator of the measured quadrature angle θ. Knowing the field amplitudes \(| {\overline{a}}_{\hom }| ,| {\overline{a}}_{{\rm{sig}}}| \) and that \(\overline{\varDelta }=-\kappa /(2\sqrt{3})\), we can reconstruct the measured quadrature angle as the one satisfying the cancellation condition.

    A detailed characterization of the nonlinear mixing noise and an analysis of single-detector homodyne efficiency can be found in the Supplementary Information.

    Multimode Kalman filter

    The continuous position measurement of an oscillator at frequency Ωm can be viewed as a form of heterodyne measurement of two orthogonal mechanical quadratures of motion \(\widehat{X}\) and \(\widehat{Y}\) that rotate with frequency Ωm. IQ demodulation can then be carried out at the mechanical frequency Ωm. This results in two independent measurement channels of two orthogonal mechanical quadratures with independent measurement noise.

    We work in a parameter regime in which the measurement rate is significantly smaller than the frequency of the mechanical mode, such that we can perform IQ demodulation of the mechanical motion at Ωm to obtain the slowly varying \(\widehat{X},\widehat{Y}\) quadratures. Their evolution is described by decoupled quantum master equations33. In this parameter regime, only thermal coherent states are prepared through the measurement process. These states are essentially thermal states displaced from the origin of the phase space and belong to the larger group of Gaussian states.

    We operate in the fast-cavity limit Ωmκ, so the cavity dynamics are simplified in our modelling. After IQ demodulation, the normalized photocurrent signal is described by

    $${\bf{i}}(t){\rm{d}}t={\rm{d}}{\bf{W}}(t)+\sum _{i}\sqrt{4{\varGamma }_{{\rm{meas}}}^{i}}\langle {\widehat{{\bf{r}}}}_{i}\rangle (t){\rm{d}}t$$

    (1)

    where the subscript i denotes different mechanical modes, \({\bf{i}}=\left[\begin{array}{c}{i}_{X}\\ {i}_{Y}\end{array}\right]\), \({\widehat{{\bf{r}}}}_{i}=\left[\begin{array}{c}{\widehat{X}}_{i}\\ {\widehat{Y}}_{i}\end{array}\right]\) and \({\rm{d}}{\bf{W}}=\left[\begin{array}{c}{\rm{d}}{W}_{X}\\ {\rm{d}}{W}_{Y}\end{array}\right]\). The Wiener increment dWX,Y(t) = ξ(t)dt is defined in terms of an ideal unit Gaussian white noise process \(\langle \xi (t)\xi ({t}^{{\prime} })\rangle =\delta (t-{t}^{{\prime} })\).

    As the measurement is purely linear, the system remains in a Gaussian state54, and the dynamics are completely captured by the expectation values of the quadratures Xi, Yi and their covariance matrix C. We derive the time evolution of the quadrature expectation values as

    $${\rm{d}}\langle {\widehat{{\bf{r}}}}_{i}\rangle ={A}_{i}\langle {\widehat{{\bf{r}}}}_{i}\rangle {\rm{d}}t+2{B}_{i}{\rm{d}}{\bf{W}}(t),$$

    (2)

    where

    $${A}_{i}=\left[\begin{array}{cc}-{\varGamma }_{{\rm{m}}}^{i}\,/2 & {\varOmega }_{i}-{\varOmega }_{{\rm{m}}}\\ {\varOmega }_{{\rm{m}}}-{\varOmega }_{i} & -{\varGamma }_{{\rm{m}}}^{i}\,/2\end{array}\right]$$

    and

    $${B}_{i}=\left[\begin{array}{cc}{\sum }_{j}\sqrt{{\varGamma }_{{\rm{meas}}}^{j}}{C}_{{\widehat{X}}_{i}{\widehat{X}}_{j}} & {\sum }_{j}\sqrt{{\varGamma }_{{\rm{meas}}}^{j}}{C}_{{\widehat{X}}_{i}{\widehat{Y}}_{j}}\\ {\sum }_{j}\sqrt{{\varGamma }_{{\rm{meas}}}^{j}}{C}_{{\widehat{Y}}_{i}{\widehat{X}}_{j}} & {\sum }_{j}\sqrt{{\varGamma }_{{\rm{meas}}}^{j}}{C}_{{\widehat{Y}}_{i}{\widehat{Y}}_{j}}\end{array}\right].$$

    The covariance matrix elements \({C}_{\widehat{M}\widehat{N}}=\langle \widehat{M}\widehat{N}+\widehat{N}\widehat{M}\rangle /2-\langle \widehat{M}\rangle \langle \widehat{N}\rangle \) evolve as

    $$\begin{array}{l}{\dot{C}}_{{\widehat{M}}_{i}{\widehat{N}}_{j}}=-\frac{{\varGamma }_{{\rm{m}}}^{i}+{\varGamma }_{{\rm{m}}}^{j}}{2}{\dot{C}}_{{\widehat{M}}_{i}{\widehat{N}}_{j}}+{\delta }_{{\widehat{M}}_{i},{\widehat{N}}_{j}}{\varGamma }_{{\rm{th}}}^{i}+{\delta }_{M,N}\sqrt{{\varGamma }_{{\rm{qba}}}^{i}{\varGamma }_{{\rm{qba}}}^{j}}\\ \,\,+{(-1)}^{{\delta }_{M,Y}}({\varOmega }_{i}-{\varOmega }_{{\rm{m}}}){C}_{{\widehat{{\mathcal{M}}}}_{i}{\widehat{N}}_{j}}+{(-1)}^{{\delta }_{N,Y}}({\varOmega }_{j}-{\varOmega }_{{\rm{m}}}){C}_{{\widehat{M}}_{i}{\widehat{{\mathcal{N}}}}_{j}}\\ \,-4\left(\sum _{k}\sqrt{{\varGamma }_{{\rm{meas}}}^{k}}{C}_{{\widehat{M}}_{i}{\widehat{X}}_{k}}\right)\left(\sum _{l}\sqrt{{\varGamma }_{{\rm{meas}}}^{l}}{C}_{{\widehat{N}}_{j}{\widehat{X}}_{l}}\right)\\ \,-4\left(\sum _{k}\sqrt{{\varGamma }_{{\rm{meas}}}^{k}}{C}_{{\widehat{M}}_{i}{\widehat{Y}}_{k}}\right)\left(\sum _{l}\sqrt{{\varGamma }_{{\rm{meas}}}^{l}}{C}_{{\widehat{N}}_{j}{\widehat{Y}}_{l}}\right),\end{array}$$

    (3)

    where \(\widehat{{\mathcal{M}}}\) and \(\widehat{{\mathcal{N}}}\) are the canonical conjugate observables of \(\widehat{M}\) and \(\widehat{N}\).

    Equations (1)–(3) form a closed set of update equations given the measurement record i(t), and enable quadrature estimations of an arbitrary number of modes and their correlations. The thermal occupancy \({\bar{n}}_{{\rm{c}}{\rm{o}}{\rm{n}}{\rm{d}},i}\) of a specific mechanical mode is determined by the quadrature phase-space variances \({V}_{{\widehat{X}}_{i}}={C}_{{\widehat{X}}_{i}{\widehat{X}}_{i}}\) and \({V}_{{\widehat{Y}}_{i}}={C}_{{\widehat{Y}}_{i}{\widehat{Y}}_{i}}\), which are both equal to \({\bar{n}}_{{\rm{c}}{\rm{o}}{\rm{n}}{\rm{d}},i}+1/2\).

    We record the voltage output from the photodetector using an UHFLI lock-in amplifier (Zurich Instruments), digitizing the signal at a 14-MHz sampling rate for a total duration of 2 s, and we store the data digitally for post-processing. The noise power spectrum density of the digitized signal is compared with that simultaneously measured on a real-time spectrum analyser, to rule out signal-to-noise ratio degradation from the digitization noise. Details of an additional filtering step are discussed in the Supplementary Information. After filtering, only the 10 mechanical modes around the defect mode frequency Ωm are kept for the multimode state estimation study.

    To perform the multimode state estimation, we extract the required system parameters of the nearest 10 mechanical modes around Ωm by fitting the measured spectral noise density. We demodulate the signal at Ωm and feed the time-series signal i(t) to the discretized version of the update equation (2),

    $$\Delta \langle {\widehat{{\bf{r}}}}_{i}\rangle ={A}_{i}^{{\prime} }\langle {\widehat{{\bf{r}}}}_{i}\rangle \Delta t+2{B}_{i}\Delta {\bf{W}}(t)$$

    (4)

    to track all the 20 quadrature expectations at different times. Here, \({A}_{i}^{{\prime} }=\left[\begin{array}{cc}-{\varGamma }_{{\rm{m}}}^{{\prime} i}\,/2 & {\varOmega }_{i}^{{\prime} }-{\varOmega }_{{\rm{m}}}\\ {\varOmega }_{{\rm{m}}}-{\varOmega }_{i}^{{\prime} } & -{\varGamma }_{{\rm{m}}}^{{\prime} i}\,/2\end{array}\right]\) contains modified mechanical parameters:

    $$\begin{array}{l}{\varGamma }_{{\rm{m}}}^{{\prime} i}={\varGamma }_{{\rm{m}}}^{i}+2{\rm{Re}}\,\left[-\frac{1-\cos (({\varOmega }_{i}-{\varOmega }_{{\rm{m}}})\Delta t)}{\Delta t}\right]\\ {\varOmega }_{i}^{{\prime} }={\varOmega }_{i}-{\rm{Im}}\,\left[i({\varOmega }_{i}-{\varOmega }_{{\rm{m}}})-\frac{{{\rm{e}}}^{{\rm{i}}({\varOmega }_{i}-{\varOmega }_{{\rm{m}}})\Delta t}-1}{\Delta t}\right]\end{array}$$

    to compensate for the influence of discretization on the state estimation performance compared with an ideal continuous one.

    The evolution of the matrix Bi, involving 210 independent covariance matrix elements, can be computed independently from the sampled time-domain data. Therefore, we calculate it following equation (3), with an update rate of 140 MHz to mitigate the discretization effect, which is then used for the update equation (4) at the sampling rate of 14 MHz. The verification of the correct implementation of the multimode Kalman filter is shown in the Supplementary Information.

    To experimentally reconstruct the covariance matrix from the estimated quadrature data, we use the retrodiction method. The retrodiction method uses the measurement record in the future as a separate state estimation result. We derived the retrodiction update equations39 and found that they are identical to the prediction update equations, except with negative mechanical frequencies. As a result, we have the following relations between covariance matrix elements estimated by prediction and retrodiction (respectively identified by the superscripts p and r):

    $$\begin{array}{l}{C}_{{\widehat{X}}_{i}{\widehat{X}}_{j}}^{{\rm{p}}}={C}_{{\widehat{X}}_{i}{\widehat{X}}_{j}}^{{\rm{r}}}\\ \,{C}_{{\widehat{Y}}_{i}{\widehat{Y}}_{j}}^{{\rm{p}}}={C}_{{\widehat{Y}}_{i}{\widehat{Y}}_{j}}^{{\rm{r}}}\\ {C}_{{\widehat{X}}_{i}{\widehat{Y}}_{j}}^{{\rm{p}}}=-{C}_{{\widehat{X}}_{i}{\widehat{Y}}_{j}}^{{\rm{r}}}.\end{array}$$

    For each time trace slice (1 ms), we calculate the difference between the prediction and retrodiction results \({\langle \widehat{{\bf{r}}}\rangle }_{{\rm{r}}}-{\langle \widehat{{\bf{r}}}\rangle }_{{\rm{p}}}\), and calculate the covariance matrix as

    $$C=\frac{1}{2}\langle \langle \left({\langle \widehat{{\bf{r}}}\rangle }_{{\rm{r}}}-{\langle \widehat{{\bf{r}}}\rangle }_{{\rm{p}}}\right)\cdot {\left({\langle \widehat{{\bf{r}}}\rangle }_{{\rm{r}}}-{\langle \widehat{{\bf{r}}}\rangle }_{{\rm{p}}}\right)}^{\top }\rangle \rangle $$

    where is the statistical average over all the time trace slices, and \(\widehat{{\bf{r}}}=\left[\cdots ,{\widehat{X}}_{i},{\widehat{Y}}_{i},\cdots \right]\). The symbolT indicates the transposed vector.

    For a system consisting of several mechanical modes that are not sufficiently separated in frequency (Ωi − Ωj not significantly faster than any other rates in the system), cross-correlations between different mechanical modes emerge because of common measurement imprecision noise and common quantum backaction force. This generally leads to higher quadrature variance because of the effectively reduced measurement efficiency of individual modes. To decouple the mechanical oscillators that are interacting because of the spectral overlap and the measurement process, we define a new set of collective motional modes through a symplectic (canonical) transformation of quadrature basis U that diagonalizes the covariance matrix UCU = V (ref. 55). As the covariance matrix is real and symmetric, the elements of U are always real, which is required for real observables. The transformation can be understood as a normal mode decomposition of the collective Gaussian state that preserves the commutation relations, as opposed to conventional diagonalization using unitary matrices. This is represented by the requirement of the symplectic transformation UΩU = Ω, where \(\varOmega =\left[\begin{array}{cc}0 & {I}_{N}\\ -{I}_{N} & 0\end{array}\right]\) is the N-mode symplectic form and IN is the N × N identity matrix. We find that in the new quadrature basis based on the diagonalized covariance matrix, the defect mode is only weakly modified. The transformation coefficients for the defect mode are shown in the Supplementary Information.

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  • Observation of plaid-like spin splitting in a noncoplanar antiferromagnet

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  • Autonomous transposons tune their sequences to ensure somatic suppression

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    Cell culture and generation of stable cell lines

    Flp-In T-REx HEK293 (Thermo Fisher Scientific, catalogue no. R78007) cells were maintained according to the manufacturer’s recommendations. Cells were cultured in DMEM with glutamax supplemented by Na-Pyruvate and High Glucose (Thermo Fisher Scientific, catalogue no. 31966-021) in the presence of 10% fetal bovine serum (FBS; Thermo Fisher Scientific, catalogue no. 10270106) and penicillin/streptomycin (Thermo Fisher Scientific, catalogue no. 15140-122). Before their introduction, transgene cells were cultured at a final concentration of 100 µg ml−1 zeocin (Thermo Fisher Scientific, catalogue no. R250-01) and 15 µg ml−1 blasticidin (Thermo Fisher Scientific, catalogue no. A1113903). For generation of stable cell lines, pOG44 (Thermo Fisher Scientific, catalogue no. V600520) was cotransfected with pcDNA5/FRT/TO (Thermo Fisher Scientific, catalogue no. V652020) containing the gene of interest at a 9:1 ratio. Cells were transfected with Lipofectamine 2000 (Thermo Fisher Scientific, catalogue no. 11668019) on a six-well-plate format with 1 µg of DNA (that is, 900 ng of pOG44 and 100 ng of pcDNA5/FRT/TO + GOI) according to the transfection protocol provided by the manufacturer. All transgenes were cloned with an N-terminal His6-biotinylation sequence-His6 tandem (HBH) tag that allows rapid and ultraclean purification without the use of antibodies. We also added a 3× FLAG tag immediately before the HBH tag to increase the versatility of the construct, which we refer to as the 3FHBH tag. Twenty-four hours following transfection, cells were split among three wells of a six-well plate at dilution ratios of 1:6, 2:6 and 3:6 to allow efficient selection of hygromycin B (Thermo Fisher Scientific, catalogue no. 10687010). Hygromycin selection was started 48 h following the transfection time point, with a final concentration of 150 µg ml−1, and refreshed every 3–4 days until control, non-transfected cells on a separate plate were totally dead. Induction of the transgene was performed overnight at a final concentration of 0.1 µg ml−1 doxycycline (DOX). Cells were validated by immunoblotting of whole-cell lysates.

    An endogenous biotin acceptor peptide affinity tag and a FLAG tag were inserted into the Safb gene locus for mouse and fly cell lines using CRISPaint. The mouse Flp-In 3T3 cell line was purchased from Thermo Fisher Scientific (catalogue no. R76107) and cultured according to the manufacturer’s instructions. Vells were cultured in DMEM (Thermo Fisher Scientific, catalogue no. 31966-021) in the presence of 10% FBS (Thermo Fisher Scientific, catalogue no. 10270106) and penicillin/streptomycin (Thermo Fisher Scientific, catalogue no. 15140-122). The Drosophila S2R+ -MT::Cas9 cell line was purchased from DGRC (DGRC stock no. 268) and cultured in S2 medium (Thermo Fisher Scientific, catalogue no. 21720024) in the presence of 10% FBS (Thermo Fisher Scientific, catalogue no. 10270106). For CRISPaint56 constructs (see Supplementary Table 2 for a list of single-guide RNAs), cells were cotransfected with three plasmids according to the CRISPaint protocol on the six-well-plate format using FuGene HD (Promega, catalogue no. E2311). Twenty-four hours following transfection, cells were expanded on 10 cm culture plates to facilitate efficient selection of puromycin (Thermo Fisher Scientific, catalogue no. A1113803). Puromycin selection is provided in the tag construct and is driven by expression from the gene locus (in this case, either the mouse or fly Safb1 gene locus). Puromycin selection was started 48 h following transfection, at 1 µg ml−1 final concentration, and was refreshed every 2 days and, in total, was maintained until all untransfected 3T3 or S2 cells were dead. Cells were validated by immunoblotting of whole-cell lysates.

    The HeLa cell line (ACC57) was purchased from Deutsche Sammlung von Mikroorganismen und Zellkulturen and maintained in the same medium as the Flp-In 3T3 cell line, but with the addition of non-essential amino acids (Thermo Fisher Scientific, catalogue no. 11140050).

    Mouse N2A cells were maintained in DMEM, and stably expressing 3× FLAG-Cas9 or 3× FLAG-SAFB1 (Extended Data Fig. 10g) was created by cotransfection of cells with plasmids expressing the protein of interest (Cas9, SAFB1 or control) under the EF1alpha promoter flanked by PiggyBack inverted repeats, together with a plasmid expressing PiggyBac transposase. In this design, because neomycin resistance was coupled to transgene expression via an IRES element, cells were selected with 1 mg ml−1 geneticin until none remained in control transfected cells.

    Cell lines (human Flp-In T-REx HEK293, human HeLa, human HCT116, mouse Flp-In 3T3, mouse N2A and fly S2R+) were all purchased from vendors or repositories or provided by colleagues (as described above), and no further authentication of cell lines was performed following purchase. Routine mycoplasma contamination tests were performed on all cell lines using the Jena Biosciences Mycoplasma (PCR-based) detection kit (Jena Biosciences, no. PP-401).

    FLASH

    Cells on 15 cm dishes were washed with 6 ml of ice-cold PBS and UV-crosslinked with 0.199 mJ cm2 UV-C light, after which they were pelleted, snap-frozen in liquid nitrogen and stored at −80 °C until use. Pellets were resuspended in 600 µl of 1× native lysis buffer (NLB) with protease inhibitors and briefly sonicated in a Bioruptor water bath sonicator (30 s on, 30 s off, five cycles at 4 °C). Lysates were then centrifuged at 20,000 relative centrifugal force (rcf) for 10 min at 4 °C to remove insoluble material. Supernatant was transferred to a fresh tube with 25 µl of MyONE C1 streptavidin beads (Thermo) and incubated in a cold room with end-to-end rotation for 1 h. Beads were washed once with high-salt buffer (HSB), once with non-denaturing buffer (NDB), treated with 0.02 U µl−1 RNase I (Thermo) in 100 µl of NDB for 3 min at 37 °C and immediately placed on ice to stop the reaction. Beads were then washed once each with HSB and NDB. RNA ends were repaired with T4 polynucleotide kinase, after which barcoded s-oligos were ligated with T4 RNA ligase 1 for 90 min at 25 °C. The 3′ phosphate at the 3′ end of each s-oligo was removed with recombinant shrimp alkaline phosphatase (NEB, M0371) and beads were washed once each with lithium dodecyl sulfate buffer, protein lysis buffer and HSB, and finally with NDB. RNA was released by treatment with proteinase K and purified using Oligo Clean and Concentrator columns (Zymo). Reverse transcription was carried out with SuperScript III and samples then treated with RNase H (NEB) to phosphorylate the 5′-end of the cDNA molecule. Following a final round of purification with Oligo Clean and Concentrator columns (Zymo), cDNA was circularized with CircLigaseII (Lucigen) and amplified with Q5 polymerase (NEB). PCR products were purified with solid-phase reversible immobilization beads, quality controlled with Bioanalyzer and subjected to high-throughput sequencing.

    FLASH data processing

    Paired-end reads were merged with bbmerge.sh v. 38.72 using the following command: bbmerge.sh in1 = {R1.fastq.gz} in2 = {R2.fastq.gz} out = {merged.fastq.gz} outu1 = {unmerged.R1.fastq.gz} outu2 = {unmerged.R2.fastq.gz} ihist = {histogram.txt} adapter1=AGATCGGAAGAGCACACGTCTGAACTCCAGTCACCCAACAATCTC adapter2=AGATCGGAAGAGCGTCGTGTAGGGAAAGAGTGTAGATCTCGGTGG –mininsert=1. Short inserts (below 20 nt, following removal of the unique molecular identifier (UMI) and internal index) were removed with bbduk.sh v. 38.72 bbduk.sh in = {infile} out = {out} minlen=34. The UMI was removed from reads and written to the header with UMI_tools v.1.0.0: umi_tools extract –bc-pattern=NNNXXXXXXNNNNN -I {IN.fastq.gz} -−3prime –stdout = {OUT.fastq.gz}, followed by separation of replicates with flexbar v.3.5.0: flexbar -r INPUT.fastq.gz -b barcodes.fa –barcode-trim-end RTAIL –barcode-error-rate 0.2 –zip-output GZ. Reads were aligned first to abundant RNAs such as transfer RNA, small nuclear RNA, small nucleolar RNA and ribonuclear RNA, then to the genome with bowtie2 v.2.3.5: bowtie2 –no-unal –un-gz -L 16 –very-sensitive-local -x bt2_index -U fastq_in.fastq.gz -o bam_out.bam. Unaligned reads were remapped to the genome with bbmap.sh v.38.72 to capture spliced reads: bbmap.sh -Xmx50G in = {fastq_in} out = {bam_out} outu = {unmapped_out} ref = {reference.fa} sam=1.3 mappedonly=t mdtag=t trimreaddescriptions=t nodisk. Finally, PCR duplicates were removed using UMI-tools: umi_tools dedup -I in_bam -S out_bam –spliced-is-unique –soft-clip-threshold 3 –output-stats = {stats}. Coverage files were generated with bamCoverage v.3.3.1: bamCoverage -b bam –filterRNAstrand [forward | reverse] –binSize 1 –normalizeUsing CPM –exactScaling -o out_file.

    UMAP of FLASH data

    For construction of the UMAP, peak calling was carried out on all profiles using HOMER: findPeaks {tag_directory} -style factor -strand separate -o {peaks.txt} -i {background_tag_directory}. Peaks from all profiles were then merged with: mergePeaks -strand -d given -matrix {peaks1.txt peaks2.txt …} > merged.peaks.txt. A count matrix, using all alignments from all profiles against merged peaks, was then created with featureCounts v.2.0.1: featureCounts -F SAF -Q 10 –primary -s 1 -T 12 -a {merged_peaks} -o {merged_peaks.counts.txt} {all_bam_files}. The count matrix was imported into a Jupyter notebook with pandas: peaks = pd.read_csv(“merged_peaks.counts.txt”, sep = ”\t”, index_col = ”Geneid”), scaled with sklearn.preprocessing.StardardScaler: peaks_scaled = StandardScaler().fit_transform(peaks), which was then used to create the UMAP: peaks_scaled_mapper = umap.UMAP(n_neighbors=15, random_state=42).fit(peaks_scaled), and plotted using umap.plot.points function. Clusters were called with HDBSCAN: clusterable_embedding = umap.UMAP(n_neighbors=30, min_dist=0.0, n_components=14, random_state=42).fit_transform(peaks_scaled), then hdbscan_labels = hdbscan.HDBSCAN(min_samples=100, min_cluster_size=600, core_dist_n_jobs=1).fit_predict(clusterable_embedding).

    Sample and library preparation for RNA-seq

    Flp-In T-REx HEK293 and HeLa ACC57 cells were transfected at a final concentration of 5 nM each (in the case of triple knockdown, total siRNA concentration became 15 nM and hence single-knockdown transfections were increased to 15 nM with the addition of 10 nM negative control siRNA) using Silencer Select siRNAs (Thermo Fisher Scientific, catalogue no. 4427037 for 1 nM scale) and RNAiMAX (Thermo Fisher Scientific, catalogue no. 13778030) on six-well plates (around 200,000 were used per replicate). Silencer Select siRNAs are 21 nt long, chemically modified (the exact modification is proprietary; Thermo Fisher) and reduce overall off-target effects by up to 90% without compromising potency. This modification also exaggerates strand bias, which correlates with better knockdown, and therefore they are 5- to 100-fold more potent than other siRNAs. The siRNA ID for human SAFB1 is s12452, for SAFB2 is s18599 and for SLTM is s36384. Cells were harvested on the second day of knockdown.

    The Silencer Select siRNAs used were s29362 for MPP8 was s23449 for TASOR.

    Flp-In 3T3 cells were first reverse transfected (roughly 100,000 per replicate) with 5 nM siRNA, boosted with the same amount 24 h following knockdown (forward transfected) and harvested on the third day following initial transfection. The siRNA ID for mouse Safb1 is s104978, for Safb2 is s104977 and, because the human SLTM siRNA also targets mouse mRNA, the same siRNA was used.

    Drosophila S2R+ cells (DGRC no. 150) were transfected with control dsRNA against GFP or Saf-B using FuGENE HD (Promega) for 3 days, after which cells were harvested for RNA isolation.

    Total RNA from human, mouse or Drosophila cells was extracted with the Quick-RNA MicroPrep kit (Zymo). Polyadenylated RNA was isolated from total RNA with the Dynabeads mRNA Purification Kit (Thermo). Purification was carried out twice to enrich poly(A)+ RNA. Sequencing libraries were generated using the KAPA Stranded RNA-Seq Library Preparation Kit (Roche).

    Isolation of nuclear and cytoplasmic RNA for RNA-seq

    Forty-eight hours following siRNA transfection (control or SAFB1 + SAFB2 + SLTM, 5 nM each), approximately 1 million Flp-ln T-REx HEK293 cells per replicate were trypsinized and either used directly for RNA isolation (total sample) or resuspended with a buffer containing 0.5% Igepal CA-630 to separate nuclear and cytoplasmic fractions, as described in ref. 57. Nuclear and cytoplasmic RNAs were isolated with the Quick-RNA MicroPrep kit (Zymo). Ribo-depleted RNA-seq samples were prepared using the KAPA RNA HyperPrep Kit with RiboErase (HMR) (no. KK8560, Roche).

    Transient transfections in rescue experiments and sample preparation for qPCR detection

    SAFB triple knockdown was performed on Flp-In T-REx HEK293 cells as described above, and then FuGENE HD forward transfected with WT or truncation mutants as shown in Extended Data Fig. 7f while at the same time refreshing the medium 6 h following transfection of siRNAs. Transgenes were induced on day 1 of knockdown with 0.1 µg ml−1 DOX for 24 h. On day 2 of knockdown, total RNA extracts were prepared with the Zymo Quick-RNA Kit and first-strand cDNA synthesis was carried out with PrimerScript RT Master Mix (TaKaRa, no. RR036A). Quantitative real-time PCR was performed using the oligos listed in Supplementary Table 1 with the Blue S’Green qPCR Kit (Biozym, no. 331416).

    ONT direct RNA-seq

    Isolation of polyA-enriched mRNA from Flp-ln T-REx HEK293 cells treated with either control siRNA or siRNAs against SAFB1, SAFB2 and SLTM (5 nM each) for 2 days was carried out using the Dynabeads mRNA DIRECT purification kit (Thermo Fisher Scientific) following the manufacturer’s instructions, with minor modifications. In brief, approximately 4 × 106 cells were subjected to the standard protocol and hybridization of the beads/mRNA complex was carried out for 10 min on a Mini Rotator (Grant-bio). DNA containing supernatant was removed and the beads were resuspended with 2 × 2 ml of buffer A following a second wash step with 2 × 1 ml of buffer B. Purified RNA was eluted with 10 µl of preheated elution buffer (10 mM Tris-HCl pH 7.5) for 5 min at 80 °C. Quantification of isolated mRNA was performed using a Qubit Fluorometer together with the RNA HS Assay kit (Thermo Fisher Scientific). For direct RNA-seq, 700 ng of freshly isolated polyA-enriched mRNA was processed according to the manufacturer’s protocol (no. SQK-RNA002). Final sequencing libraries were then loaded on R9.4 flow cells and sequenced on MinION and PromethION sequencers.

    Retrotransposition assay

    The transfection and experimental timeline for the retrotransposition assay was followed as described in ref. 18. Initially around 200,000 HeLa cells were transfected, with the same siRNAs and under the conditions listed above, on a six-well plate with 5 nM final concentration each of negative control, SAFB1, SAFB2 and SLTM siRNAs. The following day, knockdown HeLa cells were transfected with 200 ng of plasmids pYX015 (based on JM111, which has a point mutation in ORF1p) for background control and pYX017 (pCAG-driven L1RP) for L1 activity in triplicates, using Lipofectamine 2000 on a 48-well plate in triplicate. Twenty-four hours following reporter construct transfection, 2.5 µg ml−1 puromycin selection was started and maintained for 3 days (that is, day 5 of knockdown). Cells were washed with PBS before lysing with 40 µl of passive lysis buffer from the Dual-Luciferase Reporter Assay System (Promega, catalogue no. E1960). Half of the lysate was transferred to a 96-well, reading-compatible plate and measured using an Omega Lumistar machine.

    RNA–FISH

    FISH was carried out in HCT116 cells transfected with control versus siRNAs against SAFB1 and SLTM (no SAFB2 expression was detected in HCT116 cells) for 48 h using the Stellaris RNA–FISH kit (https://www.biosearchtech.com/assets/bti_stellaris_protocol_adherent_cell.pdf). Probes against L1Hs were synthesized by LGC Biosearch Technologies (see Supplementary Table 2 for sequences). Probes against GAPDH were sourced from LGC Biosearch Technologies (SMF-2026-1), provided by M. Bothe. Probes were used at a concentration of 125 nM and hybridized for 16 h at 37 °C. Samples were imaged using a Leica Stellaris 8 confocal microscope.

    EMSA with recombinant Halo, Halo-SAFB1RRM and Halo-TRA2BRRM

    The RNA-binding domain of TRA2B (residues 111:201) and SAFB1 (residues 386:485) were cloned into a plasmid encoding 10× His-TEV-Halo. Three constructs (Halo only, Halo-TRA2BRRM and Halo-SAFB1RRM) were then expressed using BL21-CodonPlus(DE3)-RIL bacteria, which were induced when an optical density of roughly 0.6 was reached, with 0.2 mM isopropyl-ß-d-thiogalactopyranoside for 4 h at 37 °C, then collected by centrifugation. Bacteria were resuspended with lysis buffer (50 mM HEPES pH 8.0, 300 mM NaCl, 5 mM imidazole and 0.05% Igepal CA-630) and disrupted with a Branson sonifier, clarified by centrifugation and filtered through a 0.45 µm membrane. Cleared lysates were incubated with cOmplete His-Tag Purification Resin (Roche), washed extensively with lysis buffer and incubated with 0.5 µM OregonGreen (Promega) on beads in lysis buffer at room temperature for 30 min for fluorescent labelling of proteins. Beads were first washed extensively with lysis buffer, then with high-salt wash buffer (50 mM Tris.Cl pH 8.0, 1 M NaCl, 5 mM imidazole) and lastly with lysis buffer. Proteins were eluted with elution buffer (50 mM Tris.Cl pH 8.0, 100 mM KCl, 200 mM imidazole). Eluates were pooled, dithiothreitol (DTT, 1 mM final concentration) and TEV protease (home-made, 6× His-tagged, approximately 1:100) were added and samples dialysed against 25 mM Tris.Cl pH 7.4, 50 mM KCl, 5% glycerol and 1 mM DTT overnight in a cold room (about 8 °C). Dialysed eluates were then incubated with cOmplete His-Tag Purification Resin (Roche) for removal of TEV protease and undigested proteins, and flowthrough was centrifuged at 23,000 rcf for 30 min and filtered through a 0.22 µm membrane to remove particulate matter. The UV spectra showed no significant absorption at 260 nm and were used to quantify purified proteins, which were then normalized and their quality checked with PAGE and Coomassie staining (Fig. 4a). Concentrations used in EMSAs were: Halo-TRA2BRRM (lanes 2–6: 3.6, 7.2, 14.4, 57.6 and 102.4 µM, respectively); Halo-SAFBRRM (lanes 7–11: 3.6, 7.2, 14.4, 57.6 and 102.4 µM, respectively); and Halo (lane 12: 102.4 µM). Lane 1 contained only those probes with no added protein.

    The RNA probes were prepared by in vitro transcription. Briefly, a plasmid containing the relevant sequence TAATACGACTCACTATAGGGAAGAAGAAGAAGAAGAAGAAGAT^ATC, in which the T7 promoter sequence is underlined, was digested with EcoRV (site of digestion, indicating that the last nucleotide of the final RNA is marked—indicated by ^), purified and in vitro transcribed using a HighYield T7 RNA Synthesis Kit (Jena Biosciences, no. RNT-101) with either 1 mM (final) CTP/UTP/GTP/ATP or 1 mM CTP/UTP/GTP and 1 mM N6-Methyl-ATP (Jena Biosciences, no. RNT-112-S), completely replacing ATP. RNA was cleaned up using SPRI beads to remove the plasmid and other potential high-molecular-weight products, then with the OCC-5 kit (Zymo). RNA was then oxidized using freshly prepared sodium periodate (250 mM in water, final concentration 10 mM; Sigma, no. 311448) in 60 mM NaOAc pH 5.5 for 1 h on ice, with tubes kept in the dark. After a further clean-up with OCC-5, RNA was then labelled with CF 647 Hydrazide (Sigma, no. SCJ4600046; 10 mM in water, 0.8 mM final concentration in approximately 120 mM NaOAc, pH 5.5) at room temperature overnight. RNA was purified with OCC-5, eluted in water and normalized to 5 µM. EMSAs were carried out in 25 mM Tris.Cl pH 7.4, 50 mM KCl, 5% glycerol and 1 mM DTT with an RNA probe of around 100 nM and the indicated concentration of the protein of interest. Following incubation of RNA and proteins on ice for 30 min, mixtures were loaded directly on a Nature 8% polyacrylamide gel cast with 0.5× Tris-borate-EDTA (final) and run in 0.5× Tris-borate-EDTA in a cold room for 45 min at 100 V (gels were prerun at 100 V for 15 min). Proteins and RNA were sequentially visualized on the same gel using a Typhoon Scanner with appropriate excitation lasers and emission filters.

    In vitro unmethylated and methylated RNA-binding assay

    Nuclei were isolated from wt-HCT116 cells using a buffer containing 0.5% Igepal CA-630, following Lubelsky and Ulitsky57, and snap-frozen in liquid nitrogen until use. Nuclei were resuspended with 500 µl of 25 mM Tris.Cl pH 7.4, 150 mM KCl, 2 mM MgCl2, 0.5% Igepal CA-630, 5% glycerol, 5 mM β-mercaptoethanol, 1× protease inhibitors and 1× PhosSTOP and sonicated with a Branson sonifier. Next, 15 µl of TURBO-DNase was added followed by incubation at 25 °C for 20 min and then by the slow addition to the lysate of 1.5 m of base buffer (25 mM Tris.Cl pH 7.4, 50 mM KCl, 5% glycerol) to bring the KCl concentration to 75 mM and Igepal CA-630 concentration to 0.125% (final). Lysate was incubated with 50 µl of Pierce Control Agarose Resin (no. 26150) for 20 min, with rotation in a cold room, and spun down at full speed for 10 min at 4 °C to remove insoluble material. A 949 bp fragment of L1 ORF2 was amplified from pYX017 using primers AATAATACGACTCACTATAGCGTATCACCACCGATCCCACAG (T7 promoter underlined) and GGCTGAGACGATGGGGTTTT and in vitro transcribed using a HighYield T7 RNA Synthesis Kit (Jena Biosciences, no. RNT-101) with either 1 mM (final) CTP/UTP/GTP/ATP or 1 mM CTP/UTP/GTP and 1 mM N6-methyl-ATP (Jena Biosciences, no. RNT-112-S), completely replacing ATP. RQ1 DNase (Promega) was added to each reaction with incubation for for 20 min at 37 °C, after which RNA was cleaned up using RCC-25 (Zymo) and oxidized with freshly prepared sodium periodate (250 mM in water, final concentration 10 mM; Sigma, no. 311448) in 60 mM NaOAc pH 5.5 for 1 h on ice, with tubes kept in the dark. After a further clean-up with RCC-25, RNA was then labelled with biotin Hydrazide (Sigma, no. 87639; 50 mM in DMSO, 2 mM final concentration in approximately 120 mM NaOAc, pH 5.5) at room temperature overnight. RNA was purified with RCC-25, eluted in water and quantified with Nanodrop, then 5 µg of each RNA or buffer was incubated with 25 µl of MyONE C1 streptavidin beads in base buffer + 0.1% Igepal CA-630 for 1 h at room temperature and washed twice with base buffer + 0.1% Igepal CA-630. The nuclear lysate was incubated with these beads for 1 h at 16 °C, with shaking at 1,100 rpm. Beads were washed and transferred from fresh tubes with base buffer + 0.1% Igepal CA-630. Proteins bound to the beads were eluted with base buffer + 0.1% Igepal CA-630 + 2 µl of RNaseA + T1 (no. EN0551, Thermo Fisher Scientific) for 30 min at 30 °C and demonstrated by immunoblotting.

    RNA blotting

    HCT116 cells were transfected with 5 nM siRNA (as indicated in Fig. 2h) then, 48 h later, were either transfected with a plasmid encoding L1Hs and driven by a minimal EF1alpha (without an intron) promoter or mock transfected. Twenty-four hours later (72 h post siRNA transfection), cells were trypsinized and resuspended with a buffer containing 0.5% Igepal CA-630, essentially as described in ref. 57. The cytoplasmic fraction was purified with RNA Clean & Concentrator columns (Zymo), 2 µg of which was loaded onto 1.2% agarose gel and electroblotted to a nylon membrane. DIG-labelled probes against ORF2 were prepared with in vitro transcription (see Supplementary Table 2 for primers) and probe hybridization, washes and imumunodetection were carried out as described in the manual of the DIG Northern Starter Kit (Roche, no. 12 039 672 910).

    p-SR (1H4) and DHX9 FLASH in SAFB-depleted cells

    Flp-In T-REx HEK293 cells were transfected with either control siRNA or siRNAs against SAFB1, SAFB2 and SLTM 48 h following transfection, then washed with PBS and UV-crosslinked with 0.2 mJ cm2 UV-C light on ice. Nuclei were isolated as described in ref. 57, resuspended in 1× NLB + 5 mM MgCl2 with protease and phosphatase inhibitors and sonicated using a Branson sonifier. Following centrifugation, to remove insoluble material the supernatant was incubated with an agarose resin (Pierce, no. 26150) for 20 min in a cold room followed by further incubation with Dynabeads Protein G beads prebound to p-SR antibody (10 µl per IP; 1H4, Santa Cruz, no. sc-13509) for 90 min in a cold room. The supernatant from 1H4 IP was used for DHX9 IP (2.5 µl per IP; abcam, no. ab26271). The FLASH protocol was identical to that described above, except that all HSB washes were replaced with NLB and s-oligos were pre-dephosphorylated to skip the recombinant shrimp alkaline phosphatase treatment that could dephosphorylate SR proteins on the beads, potentially leading to their elution.

    RIP–qPCR

    Flp-In T-REx HEK293 cells were crosslinked with 0.2% formaldeyhde for 10 min at room temperature, extensively washed with PBS, resuspended with 1× NLB and sonicated using a Branson sonifier. The lysate was centrifuged at 23,000 rcf for 10 min at 4 °C to remove insoluble material and the supernatant then incubated with an agarose resin (Pierce, no. 26150) for 30 min in a cold room. Following brief centrifugation, the supernatant was used for IP with Dynabeads Protein G beads coupled to either an antibody against SAFB1 (10 µl per IP; Santa Cruz, no. sc-393403) or control IgG (Santa Cruz, no. sc-2025) overnight in a cold room. Beads were washed with 1× NLB and bead-bound RNA was eluted with proteinase K, as described above, purified using RCC-5 (Zymo) and utilized for RT–qPCR.

    Generation of the Dnmt3c-null allele

    Dnmt3C knockout animals were generated as described in ref. 58. For specific abolition of enzymatic activity we designed a sgRNA against the methyltransferase domain of Dnmt3C targeted to exon 15 with the following protospacer sequence: 5′-GGACATCTCACGATTCCTGG-3′. P0 animals were genotyped using Sanger sequencing following PCR with primers 5′-CTGGCCGGCTCTTCTTTGAG-3′ and 5′-GGAAATCATTCCCACCTGTCAGC-3′. The founding animal was chosen based on a 31 bp deletion, which resulted not only in a frameshift mutation beginning at codon 598 but simultaneous removal of a PfoI restriction enzyme digestion site for straightforward genotyping. The founder mutation was subsequently backcrossed into the C57BL/6 J background. Homozygous knockout males were validated as infertile, with significantly smaller and disordered testes by P42, as reported previously51. The generation of these experimental animals was regulated following ethical review by Yale University Institutional Animal Care and Use Committee (protocol no. 2020-20357) and was performed according to governmental and public health service requirements. No sample size selection, randomization or blinding was performed.

    Direct antibody labelling

    The Mix-n-Stain CF488 A Antibody Labelling Kit (Biotium, no. 92253) and Mix-n-Stain CF555 Antibody Labelling Kit (Biotium, no. 92254) were used to label rabbit antihuman SAFB1/SAFB antibody (LSBio, LS-C286411) and rabbit anti-LINE-1-ORF1p antibody (abcam, no. ab216324), respectively. The standard protocol listed on the product website was followed, including the ultrafiltration protocol, with minor modifications. In brief, 25–35 μg of antibody was placed in the ultrafiltration vial provided and centrifuged at 14,000g for 2 min to remove all liquid. Depending on the initial amount of antibody, antibodies were eluted in 1× PBS to a final concentration of 0.75 ng μl−1 and the appropriate volume of 10X Mix-n-Stain Reaction Buffer added. The entire solution was transferred to the vial containing the dye and the labelling reaction allowed to proceed at room temperature (22–23 °C) in the dark for 30 min. Finally, 150 μl of storage buffer was added to each reaction with storage in aliquots of 50 μl at −20 °C until use.

    Testis sectioning and Immunofluorescence microscopy

    Testes from P25 Dnmt3C homozygous and heterozygous mutant males were dissected and embedded in O.C.T. compound (Tissue-Tek). Using cryosectioning, 8 μm sections were obtained with a Leica CM3050S and spotted onto Fisherbrand Superfrost Plus Microscope Slides (Fisher Scientific, no. 12-550-15) and stored at −80 °C until use. For immunofluorescence detection, slides were thawed at room temperature for over 10 min before fixing in 4% paraformaldehyde for 8 min. Permeabilization and blocking were performed at room temperature for 1 h with blocking buffer (5% bovine serum albumin (BSA), 0.2% Triton X-100 and PBS). Sections were incubated with directly labelled antibodies overnight at 4 °C, followed by three 5 min washes in 1× PBS and mounting with VECTASHIELD PLUS Antifade Mounting Medium and DAPI (Vector Laboratories, no. H-2000). Images were acquired using a Leica THUNDER Imaging System at ×40 magnification.

    Mass spectrometry

    Flp-In T-REx HEK293 cells stably expressing SAFB1, SAFB2 or SLTM (same cell lines used for FLASH) were induced with 0.1 µg ml−1 DOX for 16 h in triplicate, lightly crosslinked with formaldehyde (0.016% final) at room temperature for 10 min, extensively washed with PBS, resuspended with HMGT-K200 buffer (25 mM HEPES-KOH pH 7.4, 10 mM MgCl2, 10% glycerol, 0.2% Tween-20) and homogenized using a water bath sonicator. Following centrifugation, supernatants were then incubated with MyONE C1 streptavidin beads to pull down proteins of interest. Beads were washed with HMGT-K200, 20 mM Tris-Cl pH 7.4 and 1 M NaCl and finally with 20 mM Tris-Cl pH 7.4 and 50 mM NaCl, then submitted to the in-house MS-facility for further processing. Silver gel staining was performed using a SilverQuest Silver Staining Kit (Thermo Fisher Scientific, no. LC6070) for SAFB1 to ensure that conditions were sufficiently stringent in comparison with GFP pulldown (Extended Data Fig. 7b).

    On-beads digest and mass spectrometry analysis

    Twelve samples were boiled at 95 °C and 500 rpm for 10 min, followed by tryptic digest including reduction and alkylation of cysteines. The reduction was performed by the addition of tris(2-carboxyethyl)phosphine at a final concentration of 5.5 mM at 37 °C on a rocking platform (500 rpm) for 30 min. To perform alkylation, chloroacetamide was added at a final concentration of 24 mM at room temperature on a rocking platform (500 rpm) for 30 min. Proteins were then digested with 200 ng of trypsin (Roche) per sample, shaking at 800 rpm and 37 °C for 18 h. Samples were acidified by the addition of 1.3 µl of 100% formic acid (2% final concentration), centrifuged and placed on a magnetic rack. Supernatants containing the digested peptides were transferred to a new low-protein-binding tube. Peptide desalting was performed on self-packed C18 columns in a Tip. Eluates were lyophilized and reconstituted in 19 µl of 5% acetonitrile and 2% formic acid in water, briefly vortexed and sonicated in a water bath for 30 s before injection into nano-liquid chromatography–tandem mass spectrometry (nano-LC–MS/MS).

    LC–MS/MS instrument settings for shotgun proteome profiling and data analysis

    LC–MS/MS was carried out by nanoflow reverse-phase liquid chromatography (Dionex Ultimate 3000, Thermo Scientific) coupled online to a Q-Exactive HF Orbitrap mass spectrometer (Thermo Scientific), as reported previously59. In brief, LC separation was performed using a PicoFrit analytical column (75 μm internal diameter × 50 cm length, 15 µm Tip ID; New Objectives) and packed in house with 3 µm of C18 resin (Reprosil-AQ Pur, Dr Maisch). Peptides were eluted using a gradient from 3.8 to 38% solvent B in solvent A over 120 min at a flow rate of 266 nl min−1. Solvent A was 0.1% formic acid and solvent B comprised 79.9% acetonitrile, 20% H2O and 0.1% formic acid. Nanoelectrospray was generated by the application of 3.5 kV. A cycle of one full Fourier transformation scan mass spectrum (300–1,750 m/z, resolution 60,000 at m/z 200, automatic gain control target 1 × 106) was followed by 12 data-dependent MS/MS scans (resolution of 30,000, automatic gain control target 5 × 105) with a normalized collision energy of 25 eV. To avoid repeated sequencing of the same peptides, a dynamic exclusion window of 30 s was used.

    Raw MS data were processed with MaxQuant software (v.1.6.17.0) and searched against the human proteome database UniProtKB UP000005640 (containing 75,074 protein entries, released May 2020). The parameters of MaxQuant database searching were a false discovery rate of 0.01 for proteins and peptides, a minimum peptide length of seven amino acids, a first-search mass tolerance for peptides of 20 ppm and a main search tolerance of 4.5 ppm. A maximum of two missed cleavages was allowed for the tryptic digest. Cysteine carbamidomethylation was set as a fixed modification whereas N-terminal acetylation and methionine oxidation were set as variable modifications. The MaxQuant-processed output files can be found in Supplementary Table 3, showing peptide and protein identification, accession numbers, percentage sequence coverage of the protein and q-values.

    IP

    Native whole-cell extracts prepared using 0.5× NLB were incubated with ProtG Dynabeads (Life Technologies, no. 10004D) coupled to 1 μg of either SAFB antibody (‘Antibodies’) or IgG (mouse; Santa Cruz, no. sc-2025) in a cold room for 150 min. Beads were washed twice in 0.5× NLB for 5 min then once with NDB. RNase-treated samples were resuspended in 90 µl of NDB to which 10 µl of RNaseA + T1 mix (Thermo Scientific, no EN0551) was added. Samples were then incubated at 20 °C for 15 min and washed twice with 0.5× NLB. Elution from the beads was performed in 1× protein-loading dye by incubation for 5 min at 95 °C with shaking. Interaction partners were detected using the antibodies against proteins shown in Extended Data Fig. 7 (‘Antibodies’).

    Immunofluorescence

    Cells were crosslinked with 4% methanol-free formaldehyde in PBS at room temperature for 10 min, permeabilized with 0.5% Triton X for 10 min then blocked with 5% BSA in PBS for 30 min at room temperature. Primary antibodies (further details in ‘Antibodies’) were diluted in PBS with 0.1% Triton X and 1% BSA and incubated with fixed cells at 4 °C for about 16 h. Fluorescently labelled secondary antibodies with the appropriate serotype were used to demonstrate target proteins. Hoechst 33342 was used to stain DNA.

    Antibodies

    The following antibodies were used: AFB1 (Santa Cruz, no. sc-393403), SAFB2 (Santa Cruz, no. sc-514963), SAFB1/2 (HET) (human: Merck/Sigma-Aldrich, no. sc05-588; mouse: LSBio, no. LS-C2886411), SLTM (Invitrogen, no. PA5-59154), ORF1p (human: abcam, no. ab230966; mouse: abcam, no. ab216324), TASOR (Sigma-Aldrich, no. HPA006735), 1H4 (p-SR) (Merck/Sigma-Aldrich, no. MABE50), RBM12B (Bethyl, no. A305-871A-T), RBMX (Cell Signaling Technology, no. 14794 S), NCOA5 (Bethyl, no. A300-790A-T), ZNF638 (Sigma-Aldrich, no. ZRB1186), ZNF326 (Santa Cruz, no. sc-390606), TRA2B (Bethyl, no. A305-011A-M), U2AF2 (U2AF65; Santa Cruz, no. sc-53942), TUBULIN (Santa Cruz, no. sc-32293), SRRM1 (abcam, no. ab221061), SRRM2 (SC35) (Sigma-Aldrich, no. S4045), SON (Sigma-Aldrich, no. HPA023535), DHX9 (abcam, no. ab183731), U1-70K (SySy, no. 203011), PRP8 (Santa Cruz, no. sc-55533), RNAPII (Creative Biolabs, no. CBMAB-XB0938-YC), IgG normal mouse (Santa Cruz, no. sc-2025), SRSF1 (Santa Cruz, no. sc-33652), SRSF2 (abcam, no. ab204916), SRSF3 (Elabscience, no. E-AB-32966), SRSF7 (MBL, no. RN079PW), RB1 (Cell Signaling Technology, no. 9309 S), TRA2B (Santa Cruz, no. sc-166829) and YTHDC1 (Proteintech, no. 14392-1-AP).

    TE expression analysis

    RNA-seq data from human (HEK293, HeLa, HCT116), mouse (3T3) and Drosophila (S2) cells were mapped to their respective genome (hg38, mm10 and dm6, respectively) using the snakePipes non-coding-RNA-seq pipeline60. Internally this pipeline uses TEtranscripts23, which estimates both gene and TE transcript abundance in RNA-seq data and conducts differential expression analysis on the resultant count tables, which is carried out by DESeq2 (ref. 61). The outputs of this analysis can be found in Supplementary Tables 4–11.

    SAFB peak annotation and TE enrichment

    Overlapping SAFB1, SAFB2 and SLTM regions called by HOMER on FLASH data were merged using the function IRanges::reduce(), resulting in a single set of 29,806 SAFB-bound genomic intervals (SAFB peaks), 23,136 of which were located inside GENCODE-annotated genes (within-gene SAFB peaks). All GENCODE v.29 genes located on standard chromosomes were used as a control set (n = 58,721). repeatMasker annotation was downloaded from the UCSC genome browser, and the fraction of total length contributed by different transposable elements was calculated for 23,136 SAFB peaks and 58,721 GENCODE-annotated genes, separately for TEs inserted in sense and antisense orientation. Enrichment was calculated for a subset of sense and antisense TEs by dividing the TE fraction in peaks (that is, observed TE fraction) by that in whole genes (that is, fraction expected if SAFB peaks were distributed randomly on transcripts), followed by log2-transformation of values.

    Short-read RNA-seq data analysis

    Raw RNA-seq reads were subject to adaptor and quality trimming using cutadapt 4.1. Default options were used, except for -q 16 –trim-n -m 25 -a AGATCGGAAGAGC -A AGATCGGAAGAGC.

    Trimmed reads from human and mouse cell lines were mapped to human GRCh38 (HEK293, HeLa and HCT116 cell lines) and mouse GRCm38 (3T3 cell line) genomes using the STAR 2.7.9a aligner62. To improve the sensitivity of spliced read detection and quantification, mapping was done in two passes. In the first pass, all reads were mapped simultaneously to the STAR genome index built with GENCODE gene models (v.29 for human, v.19 for mouse) using default options, with the exception of –outFilterMismatchNoverReadLmax 0.05 –outSAMtype None. In the second pass, each sequenced library was mapped to a genome index with GENCODE gene models extended with new splice junctions detected in the first pass (–sjdbFileChrStartEnd pass1.SJ.out.tab). Other non-default STAR options used included –outFilterMismatchNoverReadLmax 0.05 –quantMode GeneCounts –alignIntronMax 1000000 –alignMatesGapMax 2000000 –sjdbOverhang 100 –limitSjdbInsertNsj 2000000.

    Trimmed reads from the fruitfly S2 cell line were mapped to the dm6 genome assembly using STAR 2.7.4a, and reads were counted using featureCounts (subread package v.2.0.0).

    Differential gene expression

    Differential gene expression analysis was performed using the DESeq2 package61 on reverse-stranded gene counts from the STAR alignment step. Genes with fewer than ten mapped reads were discarded; lfcThreshold = 1 and alpha = 0.05 were used for calling of differentially expressed genes, and results were shrunk using lfcShrink(…, type = “ashr”).

    Differential exon usage

    To avoid assignment of exonic reads to SAFB peaks, within-gene SAFB peak fragments or entire peaks overlapping GENCODE v.29-annotated exons were masked and ignored in exon usage analysis. The 22,129 peaks remaining (intronic SAFB peaks) were assigned to their host genes and RNA-seq reads were counted on both annotated exons and intronic SAFB peaks using the function Rsubread::featureCounts() with default arguments, except for countMultiMappingReads = FALSE, strandSpecific = 2, juncCounts = TRUE, and isPairedEnd = TRUE. Differentially expressed SAFB peaks were identified using the DEXSeq R package63 and, for each gene, the peak with the lowest DEXSeq P value was used as a reference for gene fragmentation. In total, 5,394 affected genes were fragmented into pre- and post-peak parts. Exonic read counts were aggregated separately for pre- and post-peak fragments and their differential expression measured using DESeq. Genes hosting SAFB peaks with DEXSeq Padjusted < 0.05 and log-fold change above 2 were classified as (genes with) upregulated peaks (n = 878) whereas those hosting peaks with DEXSeq Padjusted > 0.05 and log-fold change between −0.5 and 0.5 were used as the control set (n = 1,457).

    Differential splice junction usage

    The number of RNA-seq reads supporting each splice junction was counted in the second STAR alignment pass (SJ.out.tab file). Splice junctions that could not be unambiguously assigned a host gene, or that were supported by fewer than ten reads in total across all treatments and replicates in a given cell line, were ignored. Differentially used splice junctions were identified using DEXSeq, with default settings; splice junctions were treated as feature IDs and host genes as group IDs.

    Splice site strength quantification

    For each gene in the human genome, the probability of each nucleotide acting as a splice donor or acceptor was estimated using SpliceAI26, with default options. SpliceAI scores were matched to splice junctions detected and quantified by STAR.

    Splice site to TE distance measurement

    Distances between splice sites and nearest upstream or downstream TEs were calculated for a set of ten repeat families (L1, L2, Alu, SVA, ERVL, ERV1, TcMar-Tigger, MIR, Simple_repeat, hAT_Charlie) as follows: (1) all GENCODE genes were flattened using the function IRanges::reduce() in R; (2) STAR-detected splice junctions and repetitive elements outside annotated genes were dropped; and (3) for each remaining splice donor and acceptor, the distance (in nucleotides) to the nearest sense or antisense TE within the same flattened gene was measured separately for each of the ten repeat families. Donors and acceptors within TEs were assigned the distance of 0 nt.

    New splice acceptors within SAFB peaks in human tissues

    The number of reads supporting splice junctions in the GTEx consortium tissue data was extracted using the recount3 R package64. Tissues with fewer than 1 billion spliced reads were excluded from further analysis. Alternative splicing was quantified in an intron-centric manner—that is, splicing index was calculated separately for each splice donor and acceptor. We extracted all splice junctions located within an annotated human gene, with splice donor annotated in GENCODE v.29 and splice acceptor sited within a fully intronic SAFB peak (npeaks = 16,929). A further 21,693 such splice junctions were filtered for junction where the donor participated in multiple events, had a splicing index above 1% in at least one tissue and was supported by at least 500 reads in all 27 tissues (that is, used ubiquitously), resulting in a highly stringent set of of 1,104 splice junctions.

    p-SR and DHX9 FLASH analysis

    FLASH reads uniquely mapping to the hg38 genome were counted using featureCounts on two custom gene annotation reference sets. The first of these contained exons and SAFB peaks, with exons prioritized over SAFB peaks in the case of overlaps. SAFB peaks were assigned to their host genes and treated as exons for read counting. The second reference contained genes fully fragmented into exons, repetitive elements and introns, with exons prioritized over repeats and introns, and repeats prioritized over introns where their genomic coordinates were overlapping. Whereas the first reference allows for increased sensitivity when quantifying FLASH signal on known SAFB-binding regions, the latter sacrifices sensitivity (because it contains many short genomic fragments) for the power of recognizing regions of increased binding outside of SAFB peaks, or in SAFB peaks not called by the peak-calling software. DEXSeq analysis was performed separately on exon/peak and exon/repeat/intron counts. Regions with adjusted P < 0.05 were considered differentially bound.

    Alternative polyadenylation sites

    Aligned ONT direct RNA-seq performed on control and triple KD samples was screened for their end coordinates, under the assumption that these are derived from the close proximity of a polyadenylation site. Genomic coordinates of this collection of almost 1.5 million single-nucleotide-resolution read end sites were extended by 50 nt upstream and downstream, and overlapping intervals were collapsed into a total of 274,330 putative polyadenylation regions. The number of control and triple KD reads ending in each of these regions was counted and, for each gene, the fraction of ONT reads ending in each of its polyadenylation regions was calculated separately for control and triple KD libraries. Genes supported by at least 20 reads in which the contribution of at least one polyA isoform was changed by at least 20 percentage points between triple KD and control were considered differentially polyadenylated. In total, 14,148 genes (4,433 of genomic length over 50 kb) were supported by 20 or more reads, and 247 (231 longer than 50 kb) showed differential polyA site usage.

    Locus-specific L1 quantification

    Raw reads from HEK293 fractionation RNA-seq libraries were aligned to the hg38 genome using bwa aln, and alignments further processed with L1EM65, both with default options. L1EM counts from categories ‘only’, ‘3prunon’ and ‘passive_sense’ were summed. These total read counts were combined with read counts on individual genes (GENCODE v.29 annotation), and DESeq2 differential gene expression analysis was performed together on gene and L1 counts, treating L1 elements as independent genes.

    Reporting summary

    Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

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    Animals

    All experiments were performed in accordance with UK Home Office regulations and the UK Animals (Scientific Procedures) Act of 2013 under UK Home Office licences. These licences were approved by the Wellcome Sanger Institute (WSI) Animal Welfare and Ethical Review Board. Mice were maintained in a specific-pathogen-free unit under a 12 h light and 12 h dark cycle with lights off at 19:30 and no twilight period. The ambient temperature was 21 ± 2 °C, and the humidity was 55 ± 10%. Mice were housed at 3–5 mice per cage (overall dimensions of caging: 365 mm × 207 mm × 140 mm (length × width × height), floor area, 530 cm2) in individually ventilated caging (Tecniplast, Sealsafe 1284L) receiving 60 air changes per hour. In addition to Aspen bedding substrate, standard environmental enrichment of Nestlets, a cardboard tube/tunnel and wooden chew blocks were provided. Mice were given water and diet ad libitum.

    Mouse generation

    A complete list of the mouse lines used in this study is provided in the Source Data. Most mouse mutants were generated using the well-validated ‘KO-first allele’. This strategy relies on the identification of an exon common to all transcript variants, upstream of which a LacZ cassette is inserted to make a constitutive KO/gene-trap known as a tm1a allele. In contrast to the tm1a allele, tm1b creates a frameshift mutation after Cre-mediated deletion of the loxP-flanked exon. Other allele types are also possible and have been described previously60. Mouse production was performed as described previously61. We maintained most mutant lines (73% of the mice tested in this study) on a pure inbred C57BL/6N background, with the other lines on mixed C57BL/6 backgrounds (for example, C57BL/6N;C57BL/6BrdTyrc-Brd). For the C57BL/6N background, a core colony was established using mice from Taconic Biosciences, which was refreshed at set generational points (typically ten generations) and cryopreserved at regular intervals to avoid genetic drift. The sex and age for all mice analysed is available in the Source Data. For tumour-watch studies, mice were aged for spontaneous tumour formation until they became moribund in keeping with the above-mentioned Home Office Guidelines. To ensure compliance, mice were examined twice daily for symptoms including weight loss, poor coat condition and hunched back. Tumour histology was analysed by a consultant pathologist to confirm cancer diagnoses. Mice were assigned randomly to groups on the basis of Mendelian inheritance. 

    In vivo MN screen

    The in vivo MN screen was performed according to a previously described protocol20. The samples were analysed on the LSRFortessa or Cytomics FC500 (Becton Dickinson) system with a minimum of 100,000 events collected per sample. The gating strategy used is shown in Supplementary Fig. 1. For the analysis of MN screening data, a mixed linear effect beta regression model exploring the effect of genotype on the percentage of MN, was used. This was implemented within R (glmmTMB, v.1.0.1). In detail, a regression model was fitted using flow.cytometer as a fixed effect to account for any differences arising from the instrumentation, while assay.date was fitted as a random effect to account for the variance introduced by batch (Y ~ genotype + flow.cytometer + (1|batch). The genotype effect and associated error were estimated as a marginal mean using the emmeans package (R; v.1.4.4). The significance of the genotype effect was assessed using a likelihood ratio test. Analysis code is available at GitHub.

    High-throughput phenotypic screen

    The high-throughput phenotyping pipeline used was a series of standardized tests conducted in accordance with standard operating procedures (available at IMPReSS (https://www.mousephenotype.org/impress/index) and were performed by the Mouse Genetics Project (MGP) at the Wellcome Sanger Institute (WSI). Tests covered a broad range of biological areas, including metabolism, cardiovascular, neurological and behavioural, bone, sensory and haematological systems, and plasma chemistry. Factors predicted to affect the variables were standardized where possible. If this was not possible, measures were taken to reduce potential biases, for example, the impact of different people performing the test (known as the minimized operator), and the time of day of the test, as defined by the Mouse Experimental Design Ontology (MEDO)62. The data captured with the MEDO ontology can be found at http://www.mousephenotype.org/about-impc/arrive-guidelines. Moreover, pre-established reasons were defined for quality-control failures (for example, insufficient sample, error with equipment during test) and detailed using IMPRESS, and the data inclusion/exclusion criteria were therefore standardized. All discarded data were retained and tracked in a database to enable quality-control-failed data to be audited. Phenotyping data were collected at regular intervals on age-, sex- and strain-matched WT (control) mice. On average, at least seven homozygote mice of each sex per KO line were generated for phenotyping. If no homozygotes were obtained from ≥28 offspring from heterozygote intercrosses at postnatal day 14 (P14), the line was declared homozygous lethal. Similarly, if less than 13% of the pups resulting from heterozygote intercrosses were homozygous at P14, the line was judged as being homozygous subviable. In this event, heterozygote mice were examined in the phenotyping screen. The random allocation of mice to experimental group (WT versus KO) was driven by Mendelian inheritance. Owing to the high-throughput nature of the phenotyping screen, blinding the operators to the identity of KO lines during phenotyping was not used as the cage cards used to identify the mice included genotype information. However, in a high-throughput environment without a defined hypothesis, the potential bias is minimized. In all cases, the individual mouse was considered to be the experimental unit. Further experimental design strategies (for example, exact definition of a control animal) are defined using a standardized ontology as described previously62 and are available from the IMPC portal (http://www.mousephenotype.org/about-impc/arrive-guidelines). For a few lines, phenotyping data were also generated on a mutant of the same gene at another IMPC phenotyping centre and used to augment/enrich phenotypes from WSI. In figures that show phenotyping data, if the same phenotype was assessed by multiple assays, the most statistically robust result is shown.

    Characterization of MN gene candidates in human datasets

    MN gene candidates were mapped to orthologous genes in the human genome using ENSEMBL and integrated with GWAS data on mosaic LOY26. This was performed using PAR-LOYq calls from 205,011 male participants from the UK Biobank study27. An enrichment analysis was performed across the whole dataset to test for the over-representation of MN genes at LOY GWAS loci. To do this, we first performed MAGMA analyses (v.1.08)28 using all genomic variants within each MN gene extracting gene-level associations to the LOY phenotype. Genes were annotated on the basis of their proximity to genome-wide significant loci (P < 5 × 10−8) associated with LOY, specifically if they were 500 kb up- or downstream of the LOY gene start or end position. Second, further MAGMA analyses were performed using only those variants that were predicted to have deleterious effects (for example, non-synonymous and loss of function). Genes exhibiting an FDR-corrected MAGMA P < 0.05 were considered to be significant. Finally, for genomic loci reaching at least a suggestive level of significance in the GWAS (P < 5 × 10−5), we performed SMR and HEIDI tests (v.1.02)63 using blood gene expression level data from the eQTLGen study64 and blood protein level data from the Fenland study65. For both datasets, we considered expression of a gene to be influenced by the same genomic variation as that seen in the LOY GWAS if the FDR-corrected P value for the SMR test was P < 0.05 and the P value for the HEIDI test was P > 0.01. Human genomic variation within or around the DSCC1 gene was further studied by querying associations towards the human-equivalent phenotypic traits to those observed in Dscc1-mutant mice. Specifically, GWAS on BMD66, body mass index, number of children ever born67 and LOY26 were used to ascertain gene-level associations using all available variants within the DSCC1 gene and to perform SMR and HEIDI tests against the eQTLGen data, as described above (Supplementary Table 4). For the same four traits, exome gene-burden tests were performed using phenotypic and genetic data from the UK Biobank study. Rare exome variants (minor allele frequency < 0.1%) were identified on the basis of their predicted consequence on protein function and, using VEP68 and LOFTEE69, high-confidence protein truncation variants within DSCC1 were collapsed and tested for associations towards the four traits using BOLT-LMM70,71 (Supplementary Table 4). Finally, a phenome-wide association study for common variants within DSCC1 was performed using the Open Targets Genetics Portal72 (Supplementary Table 4).

    HREM analysis

    For analysis with HREM, embryos were collected at E14.5 and fixed in Bouin’s solution overnight. After washing in PBS, the embryos were dehydrated in a graded series of methanol. They were then infiltrated and embedded in methacrylate resin (JB4, Polysciences Europe) and stained with eosin B and acridine orange, according to previously published protocols73. The polymerized resin blocks were analysed using HREM resulting in volume datasets with isotropic voxel sizes of 2.55–3 µm. Visualization and further analysis of the HREM data were performed using Amira v.6.7.0 (Thermo Fisher Scientific) and OsiriX (v.5.6, 64 bit, Pixmeo). The embryos were staged and systematically screened for abnormalities according to a standardized protocol74,75.

    Cell lines

    MEFs were prepared from E13.5 embryos, after timed matings between Dscc1+/− mice. In brief, embryos were dissected from the decidua, mechanically disrupted and cultured in DMEM supplemented with 10% fetal bovine serum (FBS), 1.0 mM l-glutamine, 0.1 mM minimal essential medium, non-essential amino acids and penicillin–streptomycin. The initial plating was defined as passage zero, and cells were subsequently maintained on a standard protocol76. SIRT1-KO HEK293 cells were obtained from Kerafast (ENH131‐FP). Cells were grown in DMEM supplemented with 10% FBS, penicillin–streptomycin and 1% GlutaMAX. RPE-1 DSCC1Δ/flox cells were obtained from Jallepalli Laboratory43, and RPE-1 TP53-KO and TP53/DSCC1-double-KO cells were obtained from the de Lange laboratory51; these cell lines were grown in DMEM supplemented with 10% FBS and 1% GlutaMAX. iPS cells were grown in Tesr-E8 supplemented with 10 μM Y-27632 ROCK inhibitor (Stem Cell). HAP1 cells77 were cultured in Iscove’s modified Dulbecco’s medium (Invitrogen), supplemented with 10% FCS (Clontech), 1% UltraGlutamin (Lonza) and 1% penicillin–streptomycin (Invitrogen). ∆PDS5A and ∆WAPL HAP1 cells were generated using CRISPR–Cas9 as described previously78,79. The CHP-212 neuroblastoma cell line (CRL-2273) was grown in RPMI with 10% FBS.

    Modification of human iPS cells was performed according to established protocols80. In brief, the Gene Editing facility at WSI generated the DSCC1-KD BOB/iPS lines. We believe these cells to be null with just 2–3% of protein expression retained (Extended Data Fig. 7) but, nonetheless, designate this a KD allele. An asymmetrical exon within the target gene was replaced with a puromycin cassette, and a frameshift indel was introduced into the other allele. A template vector containing an EF1a-puromycin cassette was constructed for each gene, incorporating two 1.5 kb homology arms designed to align with the sequence surrounding the targeted exon. Two guide RNAs (gRNAs) were designed for each exon (Extended Data Fig. 7). The template vector (2 μg), both gRNA vectors (3 μg) and hSpCas9 (4 μg) were transfected into 2 × 106 cells using the Human Stem Cell Nucleofector Kit 2 (VPH-5022, Lonza). Subsequently, cells were seeded in 10 cm2 dishes and, after 72 h, they underwent selection with 3 μg ml−1 puromycin. Single cells were then expanded and subjected to genotyping for the verification of a frameshift indel using Sanger sequencing. The resulting KO lines were cultured in the presence of 1 μg ml−1 puromycin (ant-pr-1, InvivoGen). All of the cell lines in the research laboratories that participated in this study are routinely tested for mycoplasma and STR profiled and/or validated on the basis of the presence of unique engineered alleles as described in the Reporting Summary.

    Chromosome preparation and FISH

    Metaphase preparations were performed using a standard protocol81. For M-FISH analysis, mouse-chromosome-specific DNA libraries were provided by the Flow Cytometry Core Facility of Wellcome Sanger Institute82. To make 10 tests of M-FISH probe, 500 μl of sonicated DNA was precipitated with 100 μl mouse Cot-1 DNA (Invitrogen) and resuspended in 120 μl hybridization buffer (50% formamide, 2× saline-sodium citrate (SSC), 10% dextran sulfate, 0.5 M phosphate buffer, 1× Denhardt’s solution, pH 7.4). Metaphase preparations were dropped onto precleaned microscopy slides, and then fixed in acetone (Sigma-Aldrich) for 10 min followed by dehydration through an ethanol series (70%, 90% and 100%). Metaphase spreads on slides were denatured by immersion in an alkaline denaturation solution (0.5 M NaOH, 1.0 M NaCl) for approximately 40 s, followed by rinsing in 1 M Tris-HCl (pH 7.4) solution for 3 min, 1× PBS for 3 min and dehydration through a 70%, 90% and 100% ethanol series. The M-FISH probe (10 μl for each 22 × 22 mm hybridization area) was denatured at 65 °C for 10 min before being applied onto the denatured slides. The hybridization area was sealed with a 22 × 22 mm2 coverslip and rubber cement. Hybridization was performed in a 37 °C incubator for 40–44 h. The post-hybridization washes included a 5 min stringent wash in 0.5× SSC at 75 °C, followed by a 5 min rinse in 2× SSC containing 0.05% Tween-20 (VWR) and a 2  min rinse in 1× PBS, both at room temperature. Finally, the slides were mounted with SlowFade Diamond Antifade Mountant containing 4′,6-diamidino-2-phenylindole (DAPI, Invitrogen). Images were visualized on the Zeiss AxioImager D1 fluorescence microscope equipped with narrow band-pass filters for DAPI, DEAC, FITC, CY3, TEXAS RED and CY5 fluorescence and an ORCA-EA CCD camera (Hamamatsu). M-FISH digital images were captured using the SmartCapture software (Digital Scientific UK) and processed using the SmartType Karyotyper software (Digital Scientific). At least 20 metaphases from each sample were fully karyotyped on the basis of M-FISH and enhanced DAPI banding.

    CRISPR–Cas9 screen and sequencing

    WT and DSCC1-KD iPS cells (1 × 108) were independently infected with a human genome-wide guide RNA (gRNA) lentiviral library83 that had been recloned to swap the puromycin-resistance cassette with a neomycin-resistance cassette. Both lines were infected at a multiplicity of infection of 0.1–0.2 and a library coverage of 500× in three independent replicates, which were kept independent throughout the screen. Three days after infection, 1 mg ml−1 G418 was added to the medium and cells were cultured for an additional 10 days. When cells required passaging, a minimum of 5 × 107 cells per technical replicate was maintained at a library coverage of 500×. From each replicate, PCR was performed to amplify the gRNA region, and gRNAs were sequenced as described previously83. Single-end Illumina sequencing reads of 19 nucleotides were counted for each gRNA. To identify depleted and enriched genes in the DSCC1-KD iPS cells the software package MAGeCK84 v.0.5.6 was used. Extensive quality control of the screen was performed, and this analysis is available at the GitHub for this project (https://github.com/team113sanger/Large-scale-analysis-of-genes-that-regulate-micronucleus-formation/tree/main/CRISPR_screen_QC).

    Mini-arrayed CRISPR analyses

    The CHP-212 cell line was transduced with the lentiviral Cas9 plasmid (Addgene, 52962) and selected with 5 µg ml−1 blasticidin (Thermo Fisher Scientific, 61120) for 5 days. To test the expression and cutting efficiency of Cas9, we took transformed and untransformed cells and further transduced them with a lentiviral BFP-GFP reporter virus (Addgene 67980). After 4 days, the cells were analysed using flow cytometry (CytoFLEX, Beckman Coulter) and the cutting and transduction efficiency were determined on the basis of the ratio of BFP- and GFP-positive cells as previously described85. Notably, we confirmed that cells continued to cycle and grow throughout the experiment.

    The sgRNA-BFP plasmids were from the arrayed Sanger Institute CRISPR library (Sigma-Aldrich, HSANGERV; the sequences are provided in Supplementary Table 6). Bacteria were grown in 5 ml of LB medium overnight and DNA was extracted using a DNA purification kit (Amresco) and AcroPrep Advance 96-well filter plates (Cytavia, 8032). DNA concentrations were measured using the Quant-iT PicoGreen dsDNA Reagent (10535213). For each gene, two DNA vectors containing unique sgRNAs were mixed at equal amounts and then diluted to the same concentrations and blinded. Virus was produced by transfection of the mix of sgRNAs and the packaging plasmids psPAX (Addgene, 12260) and pMD2.G (Addgene, 12259) into HEK293FT cells. Virus was collected 3 days after transfection, and the viral titre was determined by measuring BFP expression using flow cytometry (CytoFLEX, Beckman Coulter). For the arrayed targeting screen, cells were seeded into PhenoPlate 96-well plates (Perkin Elmer, 6055302), leaving the outer wells blank. After the cells had adhered, they were transduced with lentivirus at a multiplicity of infection of >80% (each gene is targeted by two distinct gRNAs to increase the KO efficiency to >80% in our hands). Cells were allowed to recover before the addition of 2 µg ml−1 of puromycin (Santa Cruz Biotechnology, sc-108071) for 48 h. After recovery, CHP-212 cells were treated with 12.5 µM of hydroxyurea (Merck, H8627) for three cell doublings and hTERT RPE-1 cells with 50 µM for 72 h (Supplementary Table 7 (HU titration)). Next, cells were fixed with 4% PFA (Alfa Aesar, 43368) and stained with TOPRO3 (Thermo Fisher Scientific, T3605). Cells were imaged using the Operetta CLS system (Perkin-Elmer) and analysed using the Harmony software (Imaging facility, CRUK Cambridge). A prescan (×5 air objective) of each well was performed to determine 180 fields of view of each well with the ideal seeding density. These fields of view were then reimaged using a ×40 water objective. For the analysis, z planes were transformed into a maximum projection, a sliding paraboloid filter was used, and the find nuclei and find cytoplasm functions were optimized to detect our cell lines in culture. Furthermore, the find spots function was used to find MN located in the cytoplasm. Other particles were excluded on the basis of staining intensity, roundness and size. Deblinding was performed after statistical analyses.

    SIRT1 KO rescue cell viability experiment

    HEK293 cells were grown in antibiotic free medium for two passages before seeding at a density of 1 × 104 cells per well in quintuplicate into 96-well plates. Then, 24 h later, ON-TARGETplus Human DSCC1 (79095) siRNA-SMARTpool (L-014300-00-0005) at 25 nM final concentration was added to the cells, along with 0.2 ml per well of Dharmafect 2 (Dharmacon) transfection reagent in serum-free medium. The next day, complete antibiotic-free medium was added. Then, 48 h after transfection, the medium was refreshed on all wells with complete antibiotic-free medium. Three days after transfection, the cell viability was determined using the Promega Cell Titer Glo 2.0 cell viability assay. Medium was aspirated from wells and 175 μl of medium along with 25 μl Cell-Titer Glo reagent were added to each well and left to incubate for 10 min at room temperature. Medium and cell viability reagent mixture (150 μl) was transferred to black-walled, clear and flat-bottom 96-well plates for reading. Luminescence was read on the CLARIOstar microplate reader (BMG LABTECH). Cell viability was calculated by normalizing to untransfected control wells.

    DSCC1 transcript analyses

    RNA extraction was performed using the Monarch total RNA miniprep kit (New England BioLabs). RNA was converted to cDNA using the High-Capacity RNA-to-cDNA kit (Thermo Fisher Scientific) using 500 ng of total RNA. Gene expression was measured on the QuantStudio 5 qPCR System (Thermo Fisher Scientific) using TaqMan gene expression assays for human DSCC1 (Hs00900361_m1) or mouse Dscc1 (Mm01195386_m1). TaqMan Universal Master Mix II with UNG-1 was used (Thermo Fisher Scientific; 4440038). Amplification parameters were as follows: 50 °C for 2 min; 95 °C for 10 min; followed by 40 cycles of 95 °C for 15 s and 60 °C for 60 s. Relative gene expression was determined on the basis of the ΔCt values between the gene of interest and housekeeping genes GAPDH (Hs02786624_g1) and 18S rRNA (Hs03003631_g1) using the Design & Analysis v.2.6.0 software from Applied Biosystems (Thermo Fisher Scientific).

    Antibodies

    The following antibodies were used: anti-CD71-FITC (SouthernBiotech, 1720-02, 0.5 mg ml−1, 1:500)20, anti-SIRT1 (rabbit, Cell Signalling, 2496S, 1:1,000), anti-centromere (Antibodies, 15-234-0001, 1:1,000), anti-rabbit Alexa 488 (Thermo Fisher Scientific, A11034, 1:2,000), goat anti-human Alexa 647 (Thermo Fisher Scientific, A21445, 1:2,000), anti-DSCC1 (H0079075-B01P, Novus Biologicals, 1:1,000), anti-HSP90 (F-8, Santa Cruz, 1:10,000), anti-HP1γ (05-690, Millipore, 1:1,000), goat-anti-mouse-PO (DAKO, P044701, 1:2,000), anti-SMC3 (Abcam, AB 9263, 1:250), anti-SMC3 (Thermo Fisher Scientific, A300-060A, 1:1,000), anti-acetyl SMC3 mouse (Sigma-Aldrich, MABE1073, 21A7, Lys105/106, 385016, 1:1,000), anti-p53 (Cell Signaling Technology, 1C12, 2524S), anti-acetyl p53 (p53-K382Ac, Abcam, ab75754, EPR358(2) to p53 acetyl K382, 1:1,000), anti-phosphorylated-histone H2A.X (Ser139) (JBW301, Sigma-Aldrich, 05-636-I, 1:1,000), anti-β-actin (Merck, A5441, 1:10,000, 5% milk), anti-GAPDH (6C5, Abcam, ab8245, 1:1,000), anti-p21 (Abcam, ab109520, 1:1,000). Uncropped western blots are provided in the Supplementary Information.

    Immunoprecipitation

    Flash-frozen cell pellets were thawed on ice and resuspended in 1 ml cell lysis buffer (25 mM Tris-HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% NP-40, 5% glycerol) freshly supplemented with 1:100 Pierce Universal Nuclease (Thermo Fisher Scientific, 88702), 1 mM DTT (Thermo Fisher Scientific, A39255) and Halt protease (Thermo Fisher Scientific, 1860932) and phosphatase inhibitor (Roche, PhosSTOP, REF: 04906845001) and incubated on ice for 30 min. The lysis buffer was also used as wash buffer. Protein was collected by centrifugation (15,000 rcf, 10 min at 4 °C), the supernatant was transferred to a fresh tube and the pellet was discarded. The protein concentration was measured using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific, 23225) according to the manufacturer’s protocol. To start the immunoprecipitation, beads (Thermo Fisher Scientific, Immunoprecipitation Kit Dynabeads Protein A, 10006D) were conjugated to the antibody according to the manufacturers protocol. After optimization, 50 µl of beads were used to conjugate 2 µg of total antibody. The protein sample was diluted (using the lysis buffer) to 1 mg ml−1 for immunoprecipitation and 1 ml of this sample was added to 50 μl of antibody conjugated beads. The protein–bead–antibody mixture was incubated on a rotator overnight at 4 °C. The sample was placed onto a magnet and the supernatant was transferred to a new tube (this was the flow-through that was retained to assess the antibody–bead uptake). The sample was washed on a rotator three times for 10 min in 1 ml lysis buffer at room temperature. In between the second and third wash, the sample was moved to a new Eppendorf tube to eliminate any proteins stuck to the tube. A single PBS wash was performed to the sample for 5 min on a rotator at room temperature, then the sample was placed onto the magnet for the supernatant to be removed. The result was assessed using western blotting. To prepare the reagents for this, 50 μl of 2× SDS loading buffer and 5 μl of 10× reducing buffer were added to the beads. The input and the flow-through were prepared by adding the correct amount of protein, 4× SDS loading buffer, 10× reducing buffer and lysis buffer to volume. These samples were boiled at 95 °C for 5 min then loaded onto the gel (Bio-Rad, 4–12% gel) and run at 180 V for 45 min.

    Full proteome analysis

    The samples were lysed in RIPA buffer plus HaltTM protease and phosphatase inhibitor cocktail (final concentration 2×, ThermoFisher Scientific) with probe sonication and heating. Samples were then centrifuged at 13,000 rpm for 15 min to remove the pellet. Protein concentrations were measured using a Pierce BCA protein assay (Thermo Fisher Scientific). A total of 100 µg of protein per sample was taken. Proteins were reduced by addition of TCEP (Tris(2-carboxyethyl) phosphine, Sigma-Aldrich), alkylated by iodoacetamide (Sigma-Aldrich) and then purified by trichloroacetic acid precipitation. Purified proteins were digested in 100 mM TEAB by trypsin (Thermo Fisher Scientific) at 1:25 (by weight) at 37 °C for 18 h. A total of 40 or 50 µg of peptides were labelled using 0.4 mg TMT10plex (Thermo Fisher Scientific) according to the manufacturer’s instructions. The samples were mixed, dried in a SpeedVac and then fractionated on the XBridge BEH C18 column (2.1 mm inner diameter (i.d.) × 150 mm, Waters) with a gradient of 5% acetonitrile/0.1% NH4OH (pH 10) to 35% CH3CN/0.1% NH4OH in 30 min (total cycle 60 min). The flow rate was at 200 µl min−1. The peptides were reconstituted in 0.1% formic acid/H2O and analysed on the Orbitrap Fusion hybrid mass spectrometer coupled with the Ultimate 3000 RSLCnano system (both from Thermo Fisher Scientific). The samples were first loaded and desalted onto a PepMap C18 nano trap (100 µm i.d. × 20 mm, 100 Å, 5 µm; Thermo Fisher Scientific), then peptides were separated on the PepMap C18 column (75 µm i.d. × 500 mm, 2 µm; Thermo Fisher Scientific) over a linear gradient of 4–33.6% CH3CN/0.1% formic acid in 180 min, with a cycle time of 210 min and a flow rate at 300 nl min−1. The MS acquisition used MS3-level quantification with Synchronous Precursor Selection (SPS) with the top speed 3 s cycle time. In brief, the Orbitrap full MS survey scan was m/z 380–1,500 with a resolution of 120,000 at m/z 200, with AGC set at 4 × 105 and 50 ms maximum injection time. Multiply charged ions (z = 2–6) with an intensity threshold at 5,000 were fragmented in an ion trap at 35% collision energy, with AGC set at 1 × 104 and 50 ms maximum injection time, and isolation width of 0.7 Da in quadrupole. The top ten MS2 fragment ions were SPS selected with an isolation width of 0.7 Da, and fragmented in higher-energy collisionally activated dissociation (HCD) at 60% normalized collision energy (NCE), and detected in the Orbitrap to obtain reporter ion intensities at a better accuracy. The resolution was set at 60,000, and the AGC set at 6 × 104 with maximum injection time at 105 ms. The dynamic exclusion was set 60 s with a ±7 ppm exclusion window. The raw files were processed using Proteome Discoverer v.2.4 (Thermo Fisher Scientific) using the Sequest HT search engine. Spectra were searched against fasta files of reviewed UniProt Homo sapiens entries (December 2021) and an in-house contamination database. The search parameters were as follows: trypsin with 2 maximum miss-cleavage sites; mass tolerances at 30 ppm for precursor and 0.6 Da for fragment ions; dynamic modifications of deamidated (N, Q) and oxidation (M); static modifications of carbamidomethyl (C) and TMT6plex (peptide N-terminus and K). Peptides were validated by Percolator with the q value set at 0.01 (strict) and 0.05 (relaxed). The TMT10plex reporter ion quantifier included 20 ppm integration tolerance on the most confident centroid peak at the MS3 level. Only unique peptides were used for quantification. The co-isolation threshold was set at 100%. Peptides with average reported S/N > 3 were used for protein quantification, and the SPS mass matches threshold was set at 50%.

    Chromatin enrichment and MS analysis

    Flash-frozen cell pellets were thawed on ice and resuspended in nuclear-extraction buffer (15 mM Tris-HCl pH 7.5, 60 mM KCl, 15 mM NaCl, 5 mM MgCl2, 1 mM CaCl2, 250 mM sucrose, 0.3% NP-40, freshly supplemented with 1 mM DTT and Halt protease and phosphatase inhibitor (Thermo Fisher Scientific)) and incubated on ice for 5 min. Nuclei were collected by centrifugation (600 rcf, 5 min at 4 °C), washed once with nuclear-extraction buffer without NP-40, pelleted again, then resuspended in prechilled hypotonic buffer (3 mM EDTA, 0.2 mM EGTA and freshly supplemented with 1 mM DTT and Halt protease and phosphatase inhibitor) and incubated on ice for 30 min to release chromatin. Chromatin was pelleted for 5 min at 1,700 rcf at 4 °C in a cooled centrifuge and subsequently washed twice with hypotonic buffer. Chromatin pellets were solubilized using probe sonication in lysis buffer 100 mM triethylammonium bicarbonate (TEAB), 1% sodium deoxycholate (SDC), 10% isopropanol, 50 mM NaCl, 1:1,000 Pierce Universal Nuclease (Thermo Fisher Scientific) supplemented with Halt protease and phosphatase inhibitor. The protein concentration was measured using the Quick Start Bradford protein assay (BioRad) according to the manufacturer’s protocol. A total of 5 mg of protein with an equal contribution from each individual sample was reduced with 5 mM Tris-2-carboxyethyl phosphine (TCEP) for 1 h, followed by alkylation with 10 mM iodoacetamide for 30 min, then digested by adding trypsin (Pierce) at final concentration 75 ng μl−1 to each sample followed by incubation for 18 h at room temperature. For chromatin proteomics, 15 μg of protein digest was taken from each sample and labelled with TMTpro multiplexing reagents (Thermo Fisher Scientific), according to the manufacturer’s protocol. SDC was precipitated with formic acid at a final concentration of 2% (v/v) and centrifuged for 5 min at 10,0000 rpm. Supernatant containing TMTpro-labelled peptides were dried with a centrifugal vacuum concentrator. The remaining peptides were cleaned up using Pierce Peptide Desalting Spin Columns (Thermo Fisher Scientific), and then dried using a speed vacuum. Acetylated peptides were enriched with the PTMScan HS Acetyl-Lysine Motif (Ac-K) Kit (Cell Signalling Technologies, 46784) according to the manufacturer’s instructions, dried using a speed vacuum, resuspended in 100 mM TEAB and labelled with TMTpro according to the manufacturer’s protocol. Acetyl-enriched peptides were fractionated using the Pierce High pH Reversed-Phase Peptide Fractionation Kit (Thermo Fisher Scientific, 84868) according to the manufacturer’s protocol, dried using a speed vacuum and resuspended in 0.1% trifluoroacetic acid (TFA). Before MS analysis of the chromatin proteome, TMTpro-labelled peptides were fractionated with high-pH reversed-phase (RP) chromatography using the Waters XBridge C18 column (2.1 mm × 150 mm, 3.5 μm) on the Dionex UltiMate 3000 high-performance liquid chromatography (HPLC) system. Mobile phase A was 0.1% ammonium hydroxide (v/v) and mobile phase B was 100% acetonitrile and 0.1% ammonium hydroxide (v/v). Peptide separation was performed with a gradient elution of 200 μl min−1 with the following steps: isocratic for 5 min at 5% phase B, gradient for 40 min to 35% phase B, gradient to 80% phase B in 5 min, isocratic for 5 min, and re-equilibrated to 5% phase B. The fractions were collected in a 96-well plate every 42 s to a total of 65 fractions, then concatenated into 12 fractions, dried and reconstituted in 0.1% TFA. The samples were analysed using a Real Time Search-SPS-MS3 method on the Orbitrap Ascend mass spectrometer coupled to a Dionex UltiMate 3000 system. From each fraction, an estimated amount of 3 μg of peptides per fraction was injected onto a C18 trapping column (Acclaim PepMap 100, 100 μm × 2 cm, 5 μm, 100 Å) at a flow rate of 10 μl min−1. The samples were processed via a 120 min low-pH gradient elution on a nanocapillary reversed-phase column (Acclaim PepMap C18, 75 μm × 50 cm, 2 μm, 100 Å) at 50 °C. MS1 scans were collected from the 400–1,600 m/z range in the Orbitrap with the following settings: resolution, 120,000; AGC, standard; injection time, auto; and including 2–6 precursor charge states. Dynamic exclusion was set to 45 s, repeat count of 1, mass tolerance of 10 ppm and the exclude isotope option was enabled. MS2 spectra were acquired in the ion trap at Turbo scan rate, HCD collision energy was set to 32% and 35 ms maximum-injection time was allowed. MS2 scans were searched against the human canonical and isoforms database (UniProt, 16 December 2022) using the Comet search engine in real time with the following filters: tryptic peptides with maximum of 1 missed cleavages, static modifications included Cys carbamidomethylation (+57.0215) and N-terminal/Lys TMTpro (+304.2071), variable modifications Asn/Gln deamidation (+0.984) and Met oxidation (+15.9949), with maximum of variable modifications set to 2; close-out was enabled with a maximum of 4 peptides per protein. Precursors matching these criteria were selected for SPS10-MS3 scans performed at an orbitrap resolution of 45,000 with the normalized HCD collision energy set to 55%, AGC set at 200% and 200 ms maximum injection time. Acetyl-enriched peptides were analysed using an MS2-HCD method with collision energy set to 35%, AGC set at 1 × 105 and 105 ms maximum injection time. The SequestHT and Comet search engines were used to analyse the acquired spectra in Proteome Discoverer v.3.0 (Thermo Fisher Scientific) for protein identification and quantification. For analysis of the chromatin proteome, the precursor mass was set to 20 ppm and fragment mass tolerance was 0.5 Da. Spectra were searched for fully tryptic peptides with a maximum of two missed cleavages. N-terminal/Lys TMTpro and carbamidomethyl at Cys were defined as static modifications. Dynamic modifications included oxidation of Met and deamidation of Asn/Gln. For peptides enriched for acetylated lysine, the precursor mass was set to 10 ppm and the fragment mass tolerance was set to 0.02 Da. Spectra were searched for fully tryptic peptides with a maximum of three missed cleavages. N-terminal TMTpro and carbamidomethyl at Cys were defined as static modifications, while dynamic modifications included oxidation of Met, deamidation of Asn/Gln, and TMTpro or acetyl at Lys. Peptide confidence was estimated using the Percolator node. Peptide FDR was set at 1% and validation was based on q value and a target–decoy database search. Spectra were searched against reviewed UniProt human protein entries. The reporter ion quantifier node included a TMTpro quantification method with an integration window tolerance of 15 ppm and an integration method based on the most confident centroid peak at the MS3 or MS2 level. Only unique peptides were used for quantification, considering protein groups for peptide uniqueness. Peptides with an average reporter signal-to-noise ratio of >3 were used for quantification. For the chromatin proteome, the data were normalized to total loading at the proteome level, whereas, for the respective acetylome, the data were corrected for loading for acetylated peptides only. Relative abundances were calculated by dividing normalized protein/peptide abundances by the average abundance of all TMTpro channels per biological replicate.

    Immunoblotting and immunofluorescence

    Cells were scraped from dishes in 2× SDS buffer (120 mM Tris-HCl pH 6.8, 4% SDS, 20% glycerol). After total protein quantification, equal protein amounts were run on 4–12% Bis-Tris NuPAGE precast gels, transferred to nitrocellulose membrane (GE Healthcare) and immunoblotted with the indicated antibodies. For chromatin fractionation, cells were washed with cold PBS and resuspended in CSK buffer (10 mM PIPES pH 7.0, 100 mM NaCl, 300 mM sucrose, 3 mM MgCl2, protein inhibitor cocktail (Roche, EDTA-free, 1 tablet per 10 ml), EGTA-free phosphatase inhibitors (1 mM NaF, 0.7 mM β-glycerol phosphate, 0.2 mM Na3VO4, 8.4 mM Na4P2O7), 0.7% Triton X-100), incubated on ice for 30 min and centrifuged at 20,000g for 10 min at 4 °C. The supernatant (soluble fraction) was collected and maintained on ice. The pellet was washed twice with cold PBS and sonicated (four pulses of 10 s at 30% amplitude with 10 s resting on ice between cycles) in CSK buffer. The protein concentration of soluble and chromatin fractions was determined using the Bradford assay and Laemmli buffer was added to the samples. Finally, the samples were boiled, centrifuged at 16,000g for 1 min and equal amounts were loaded onto SDS–PAGE gels. For immunofluorescence studies, cells on coverslips were fixed in a formaldehyde lysis solution (4% formaldehyde, 0.5% Triton X-100, 1× PBS), washed with 1× PBS and permeabilized in 0.5% Triton X-100, 1× PBS. Blocking was performed in 1× PBS, 0.1% Triton X-100, 10% FBS for 1 h, followed by incubation with primary or secondary antibodies in the same solution. Washes were performed using 1× PBST (1× PBS, 0.1% Triton X-100). Coverslips were mounted in Vectashield Mounting Medium with DAPI (Vector Laboratories, H1200-10). Images were collected on the Leica SP8 with ×63/1.4 NA oil objectives, using the Leica Application Suite X software (LAS-X). Images were deconvolved using Huygens Professional v.19.04 software (Scientific Volume Imaging); processing and analysis were performed using ImageJ v.1.53a and Adobe Illustrator 2021. All of the images shown are the projections of z optical sections.

    SIRT1 inhibition assays

    Cells were preincubated with EX 527 (selisistat; SIRT1i; Selleckchem) resuspended in DMSO or with DMSO alone for 3 days and then seeded at a density of 2.5 × 105 cells per 10 cm dish, maintaining either SIRT1i or DMSO in the culture medium. The next day, cells were treated with tamoxifen or mock treated for 24 h. The number of living cells at each timepoint was determined after trypsinization using the Countess II machine (Life Technologies). To determine the dose of tamoxifen that resulted in full depletion of DSCC1 in the hTERT RPE-1 DSCC1Δ/floxcretam cell line, cells were grown in the presence of different concentrations of the compound. After 3 days of tamoxifen treatment (Sigma-Aldrich), cell survival was observed by staining with crystal violet (Sigma-Aldrich; 1% aqueous solution). The dose that killed all DSCC1Δ/floxcretam cells, but not WT hTERT RPE-1 control cells (100 nM), was used for subsequent experiments (Extended Data Fig. 7). To determine the dose of SIRT1i that fully inhibits SIRT1 activity in cultured hTERT RPE-1 cells, the acetylation of p53 at Lys382, a bona fide SIRT1 substrate86, was examined. Cells were grown in the presence of different concentrations of SIRT1i for 3 days (Extended Data Fig. 9). To avoid interference from other histone deacetylases 5 µM vorinostat (Sigma-Aldrich) was added to the cells 2 h before gamma irradiation (5 Gy). Then, 3 h later, the samples were collected and acetylation of p53 at Lys382 was examined using western blotting.

    SIRT1 in vitro deacetylation assay

    These experiments were performed in HDAC8-KO HAP1 cells (Horizon Discovery) and also hTERT RPE-1 cells (Extended Data Fig. 9). p53 (a known SIRT1 target) was purified 5 h after gamma irradiation (10 Gy) of hTERT RPE-1 cells that were previously treated with 10 µM selisistat and 5 µM vorinostat (SAHA; Sigma-Merk, SML0061). SMC3 was purified from exponentially growing HDAC8-KO HAP1 cells. Both proteins (p53 and SMC3) were purified by immunoprecipitation as follows: cell pellets were resuspended in lysis buffer (50 mM Tris-Cl pH 8, 150 mM NaCl, 1 mM EDTA, 0.5% igepal, complete EDTA-free protein inhibitor cocktail from Roche and phosphatase inhibitor cocktails 2 and 3 from Sigma-Aldrich) and quantified. For p53, 2 mg of protein from hTERT-RPE-1 cell lysates was incubated with 20 μl of Dynabeads (protein G) and 6 μl of anti-p53 antibodies. For SMC3, 10 mg of HAP1 cell lysate was incubated with 40 μl of Dynabeads (protein A) and 12 μg of anti-SMC3 antibodies. Both incubations were performed overnight at 4 °C. The next morning, the beads were washed four times with cold lysis buffer and twice with reaction buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM MgCl2) and resuspended in 50 μl of reaction buffer. For the deacetylation reaction, 10 μl of beads was incubated with 1 μl of human recombinant SIRT1 (Sigma-Aldrich) in a total volume of 30 μl of reaction buffer supplemented with 1.5 μM NAD+. The reactions were incubated for 3 h at 30 °C with shaking. Finally, the reactions were stopped by the addition of 10 μl of 4× Laemmli sample buffer and incubation at 95 °C for 5 min. The samples were then immunoblotted with the respective antibodies.

    siRNA experiments in RPE-1 cells

    A total of 200,000 RPE-1 DSCC1Δ/floxcretam cells were seeded per well of a six-well plate and allowed to attach overnight. The cells were then transfected with either non-targeting (referred to as SCR control) or targeting siRNA at 25 nM using 5 µl DharmaFECT 1 transfection reagent (Horizon Discovery T-2001-02) according to the manufacturer’s instructions. After 24 h, the medium was replaced in all wells and cells were treated with or without 100 nM 4-OHT. Cells were incubated for a further 5 days before collecting and cell counting by trypan blue exclusion. A list of all siRNAs used is provided in Supplementary Table 6.

    MN counting in HAP1 cells

    HAP1 cells were seeded at an equal density, grown on coverslips and transfected with siRNAs targeting luciferase or DSCC1. All siRNAs were ON-TARGETplus SMARTpools manufactured by Dharmacon and used at a final concentration of 20 μM per siRNA. Transfections were performed using Invitrogen RNAiMAX (Life Technologies) according to the manufacturer’s instructions. Transfections were repeated after 48 h. After an additional 24 h, the coverslips were fixed with freshly prepared 3.7% paraformaldehyde in PBS for 7 min at room temperature. Cells were permeabilized and stained for 10 min with 0.1% Triton X-100 in PBS, supplemented with 1 μg ml−1 DAPI at room temperature. The coverslips were washed once with PBS, and mounted onto glass slides using Prolong Gold (Invitrogen). The slides were imaged and deconvolved on the THUNDER Imager (Leica Microsystems) and analysed using ImageJ (v.2.1.0/1.53k). A cell was scored as harbouring MN when the nucleus had one or more MN in its proximity. At least 400 cells were scored per condition.

    Analysis of cohesion defects in RPE-1 TP53-KO and RPE-1 TP53/
    DSCC1-double-KO cells

    RPE-1 TP53/DSCC1-double-KO cells were generated as previously reported51 and were cultured in DMEM + 8% FCS. For analysis of cohesion defects, cells were incubated for 20 min with 200 ng ml−1 demecolcine (Sigma-Aldrich), collected, incubated for 20 min in 0.075 M KCl and fixed in 3:1 methanol:acetic acid. Cells were washed in fixative three times, dropped onto microscopy slides and stained with 5% Giemsa (Merck). For each condition, cohesion defects were counted in 50 metaphases on two coded slides.

    Statistics and reproducibility

    Statistical analyses were performed using Prism (v.9.1.0/v.10.1, GraphPad) or R (v.3/v.4.3.1). All statistical details are provided in the figure legends. All experiments were performed independently at least three times, and were replicated by independent researchers using multiple models and using blinding where possible. T-tests were unpaired. 

    Reporting summary

    Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

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  • A single-cell time-lapse of mouse prenatal development from gastrula to birth

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    Data reporting

    For newly generated mouse embryo data, no statistical methods were used to predetermine sample size. Embryos used in the experiments were randomized before sample preparation. Investigators were blinded to group allocation during sample collection and data generation and analysis. Embryo collection and sci-RNA-seq3 data generation were performed by different researchers in different locations.

    Mouse embryo collection and staging

    All animal use at The Jackson Laboratory was done in accordance with the Animal Welfare Act and the AVMA Guidelines on Euthanasia, in compliance with the ILAR Guide for Care and Use of Laboratory Animals, and with prior approval from The Jackson Laboratory Animal Care and Use Committee under protocol AUS20028.

    The details of collecting the 12 mouse embryos with somite counts ranging from 0 to 12 were described previously8. In brief, C57BL/6NJ (strain 005304) mice were obtained at The Jackson Laboratory and mice were maintained via standard husbandry procedures. Timed matings were set in the afternoon and plugs were checked the following morning. Noon of the day a plug was found was defined as E0.5. On the morning of E8.5, individual decidua were removed and placed in ice cold PBS during the collection. Individual embryos were dissected free of extraembryonic membranes, imaged, and the number of somites present were noted prior to snap freezing in liquid nitrogen (Extended Data Fig. 1a). A portion of yolk sac from each embryo was collected for sex based genotyping and samples were stored at −80 °C until further processing.

    For newly processed mouse embryos, we used a combination of staging methodologies depending on gestational age of collection (Extended Data Fig. 1b–f). To maximize temporal coherence, resolution, and accuracy, we sought to stage individual embryos based on well-defined morphological criteria, rather than by gestational day alone. Embryos collected between E8.0–E10.0 were staged based upon the number of somites counted at the time of collection and further characterized by morphological features (Extended Data Fig. 1a). For E10.25–E14.75 embryos, developmental age was determined using the embryonic mouse ontogenetic staging system (eMOSS, https://limbstaging.embl.es/), which leverages dynamic changes in hindlimb bud morphology and landmark-free based morphometry to estimate the absolute developmental stage of a sample71,72. A modified staging tool, implemented in Python and exhibiting better performance on E14.0–E15.0 samples, was used to confirm staging of samples within this window (documentation and Python scripts available at https://github.com/marcomusy/welsh_embryo_stager). To distinguish samples staged via eMOSS, these samples are prefixed with ‘mE’ to indicate morphometric embryonic day (for example, mE13.5; Extended Data Fig. 1b–f). Due to the increased complexity of limb morphology at later stages automated staging beyond E15.0 is not possible. As a consequence, collections for all remaining embryonic samples (E15.0–E18.75) was performed precisely at 00:00, 06:00, 12:00 and 18:00 on the targeted day. From close inspection of limbs in this sample set we defined additional dynamics related to digit morphogenesis that allowed further binning of samples collected on days 15 and 16 (Extended Data Fig. 1b–f). Therefore, amongst samples profiled in this study, only the E17.0–E18.75 samples were staged solely by gestational age. Finally, P0 samples were collected from litters at noon of the day of birth (parturition for C57BL/6NJ occurs between E18.75 and E19.0).

    Collection of mouse pups immediately after birth

    Samples for the validation experiment on periparturition transcriptional dynamics were collected from a plugged female that was monitored for signs of labour beginning at E18.75. Following the natural delivery of 3 pups the dam was euthanized, and following removal from the uterus and extraembryonic membranes, the remaining pups were either collected immediately or placed in a warming chamber to monitor respiratory response and collected at 20-min intervals. We collected nine new pups altogether. The first 3 pups were estimated to be between 1 h to 2 h old, although this was not precisely timed (samples 1–3 in Fig. 6c and Extended Data Fig. 12a). None of these pups had nursed at the time of collection. The next two pups were taken by C-section, decapitated and snap frozen immediately; no breaths were taken (samples 4 and 5 in Fig. 6c and Extended Data Fig. 12a). The next 4 pups were taken by C-section and used for a ‘pink up’ time course, collecting one pup every 20 min (that is, 20 min, 40 min, 60 min and 80 min; samples 6–9 in Fig. 6c and Extended Data Fig. 12a). During this time, all pups remained very active and working to establish a breathing rhythm. Pup 6 had not fully pinked up at time of collection, but pups 7–9 had. Pups 8 and 9 had visible lungs in their chest cavities at 60 min. The last pup collected at 80 min was fully pink with a reasonably stable breathing rhythm. No vocalization was heard from any pups during this collection. Of note, for additional quality control, we put nuclei from previously profiled E18.75 and P0 embryos into a small number of wells of the sci-RNA-seq3 experiment in which nuclei from this validation series were processed.

    Generating data using an optimized version of sci-RNA-seq3

    Together with E8.5 data, which has been reported previously8, a total of 15 sci-RNA-seq3 experiments were performed on a total of 75 mouse embryos. At least one sample was included for every 6-h interval from E8.0 to P0, and we also included embryos with as many specific somite counts as we could for the 0–34 somite range. Multiple samples were selected for a few timepoints (for example, two samples for E13.0) to boost cell numbers. Meanwhile, we tried to ensure that both male and female mice roughly alternated at adjacent timepoints (Extended Data Fig. 2j). A detailed summary and images of individual embryos can be found in Extended Data Fig. 1 and Supplementary Table. 1.

    To generate the dataset, we used the optimized sci-RNA-seq3 protocol3 as written, adjusting the volume and type of lysis buffer to the size of the embryos. In brief, frozen embryos were pulverized on dry ice and cells were lysed with a phosphate-based, hypotonic lysis buffer containing magnesium chloride, Igepal, diethyl pyrocarbonate as an RNase inhibitor, and either sucrose or bovine serum albumin (BSA). Lysate was passed over a 20-μm filter, and the nuclei-containing flow-through was fixed with a mixture of methanol and dithiobis (succinimidyl propionate) (DSP). Nuclei were rehydrated and washed in a sucrose/PBS/Triton X-100/magnesium chloride buffer (SPBSTM), then counted and distributed into 96-well plates for reverse transcription with indexed oligonucleotide-dT primers.

    Age-specific adaptations were as follows. E10–E13 embryos use 5 ml BSA lysis buffer, E14 embryos use 10 ml BSA lysis buffer, E15–E18 embryos use 20 ml sucrose-based lysis buffer. Each of these samples were split over 48–96 wells for reverse transcription and the first round of indexing. A newborn P0 mouse requires 40 ml of sucrose-based lysis buffer, and the lysate is divided into 4 fractions for filtration and fixing because of the amount of tissue involved. The two P0 mice were each processed as an individual experiment and were each split over 384 wells for reverse transcription.

    For the mouse samples E8.0–E9.75, we used the ‘Tiny Sci’ adaptation of the optimized sci-RNA-seq33. Frozen embryos were gently resuspended in 100 μl lysis buffer to free the nuclei, then 400 μl of dithiobis (succinimidyl propionate)-methanol fixative was added. In the same tube, fixed nuclei were rehydrated, washed and then put directly into 8–32 wells for reverse transcription.

    After reverse transcription, nuclei were pooled, washed, and redistributed into fresh 96-well plates to attach a second index sequence by ligation. Then the nuclei were pooled again, washed and redistributed into the final plates. There, the nuclei would undergo second-strand synthesis, extraction, tagmentation with Tn5 transposase and finally PCR to add the final indexes. The PCR products were pooled, size-selected, and then the library was sequenced on an Illumina NovaSeq. For some experiments, a second NovaSeq run was necessary to capture the extent of the library complexity, so we would add more sequencing reads until the PCR duplication rate met a threshold of 50% or the median UMI count per cell went over 2,500. The validation dataset (Extended Data Fig. 4a–f) generated from 8–21-somite embryos was sequenced on an Illumina NextSeq.

    Processing of sci-RNA-seq3 sequencing reads

    Data from each individual sci-RNA-seq3 experiment was processed independently. For each experiment, read alignment and gene count matrix generation was performed using the pipeline that we developed for sci-RNA-seq314 (https://github.com/JunyueC/sci-RNA-seq3_pipeline). In brief, base calls were converted to fastq format using Illumina’s bcl2fastq v2.20 and demultiplexed based on PCR i5 and i7 barcodes using maximum likelihood demultiplexing package deML73 with default settings. Demultiplexed reads were filtered based on the reverse transcription (RT) index and hairpin ligation adapter index (Levenshtein edit distance (ED) < 2, including insertions and deletions) and adapter-clipped using trim_galore v0.6.5 (https://github.com/FelixKrueger/TrimGalore) with default settings. Trimmed reads were mapped to the mouse reference genome (mm10) for mouse embryo nuclei using STAR v2.6.1d74 with default settings and gene annotations (GENCODE VM12 for mouse). Uniquely mapping reads were extracted, and duplicates were removed using the UMI sequence, RT index, ligation index and read 2 end-coordinate (that is, reads with identical UMI, RT index, ligation index and tagmentation site were considered duplicates). Finally, mapped reads were split into constituent cellular indices by further demultiplexing reads using the RT index and ligation index. To generate digital expression matrices, we calculated the number of strand-specific UMIs for each cell mapping to the exonic and intronic regions of each gene with the Python v2.7.13 HTseq package75. For multi-mapping reads (that is, those mapping to multiple genes), the read were assigned to the gene for which the distance between the mapped location and the 3′ end of that gene was smallest, except in cases where the read mapped to within 100 bp of the 3′ end of more than one gene, in which case the read was discarded. For most analyses, we included both expected-strand intronic and exonic UMIs in per-gene single-cell expression matrices. After the single-cell gene count matrix was generated, cells with low quality (UMI < 200 or detected genes <100 or unmatched_rate (proportion of reads not mapping to any exon or intron) ≥ 0.4) were filtered out. Each cell was assigned to its originating mouse embryo on the basis of the reverse transcription barcode.

    Doublet removal

    We performed three steps with the goal of exhaustively detecting and removing potential doublets. Of note, all these analyses were performed separately on data from each experiment.

    First, we used Scrublet to detect doublets directly. In this step, we first randomly split the dataset into multiple subsets (six for most of the experiments) in order to reduce the time and memory requirements. We then applied the Scrublet v0.1 pipeline76 to each subset with parameters (min_count = 3, min_cells = 3, vscore_percentile = 85, n_pc = 30, expected_doublet_rate = 0.06, sim_doublet_ratio = 2, n_neighbors = 30, scaling_method = ‘log’) for doublet score calculation. Cells with doublet scores over 0.2 were annotated as detected doublets.

    Second, we performed two rounds of clustering and used the doublet annotations to identify subclusters that are enriched in doublets. The clustering was performed based on Scanpy v.1.6.020. In brief, gene counts mapping to sex chromosomes were removed, and genes with zero counts were filtered out. Each cell was normalized by the total UMI count per cell, and the top 3,000 genes with the highest variance were selected, followed by renormalizing the gene expression matrix. The data was log-transformed after adding a pseudocount, and scaled to unit variance and zero mean. The dimensionality of the data was reduced by PCA (30 components), followed by Louvain clustering with default parameters (resolution = 1). For the Louvain clustering, we first computed a neighbourhood graph using a local neighbourhood number of 50 using scanpy.pp.neighbors. We then clustered the cells into sub-groups using the Louvain algorithm implemented by the scanpy.tl.louvain function. For each cell cluster, we applied the same strategies to identify subclusters, except that we set resolution = 3 for Louvain clustering. Subclusters with a detected doublet ratio (by Scrublet) over 15% were annotated as doublet-derived subclusters. We then removed cells which are either labelled as doublets by Scrublet or that were included in doublet-derived subclusters. Altogether, 2.7% to 16.8% of cells in each experiment were removed by this procedure.

    We found that the above Scrublet and iterative clustering-based approach has difficulty identifying doublets in clusters derived from rare cell types (for example, clusters comprising less than 1% of the total cell population), so we applied a third step to further detect and remove doublets. This step uses a different strategy to cluster and subcluster the data, and then looks for subclusters whose differentially expressed genes differ from those of their associated clusters. This step consists of a series of ten substeps. (1) We reduced each cell’s expression vector to retain only protein-coding genes, long intergenic non-coding RNAs (lincRNAs) and pseudogenes. (2) Genes expressed in fewer than 10 cells and cells in which fewer than 100 genes were detected were further filtered out. (3) The dimensionality of the data was reduced by PCA (50 components) first on the top 5,000 most highly dispersed genes and then with UMAP (max_components = 2, n_neighbors = 50, min_dist = 0.1, metric = ‘cosine’) using Monocle 3-alpha14. (4) Cell clusters were identified in UMAP 2D space using the Louvain algorithm implemented in Monocle 3-alpha (resolution = 10−6). Cell partitions were detected using the partitionCells function implemented in Monocle 3-alpha. This function applies algorithms that automatically partition cells to learn disjoint or parallel trajectories based on concepts from ‘approximate graph abstraction’77. (5) We took the cell partitions identified by Monocle 3-alpha (cell clusters were used instead for three experiments that profiled embryos before E10), downsampled each partition to 2,500 cells, and computed differentially expressed genes across cell partitions with the top_markers function of Monocle 3 (reference_cells = 1000). (6) We selected a gene set combining the top ten gene markers for each cell partition (filtering out genes with fraction_expressing <0.1 and then ordering by pseudo_R2). (7) Cells from each main cell partition were subjected to dimensionality reduction by PCA (10 components) on the selected set of top partition-specific gene markers. (8) Each cell partition was further reduced to 2D using UMAP (max_components = 2, n_neighbors = 50, min_dist = 0.1, metric = ‘cosine’). (9) The cells within each partition were further sub-clustered using the Louvain algorithm implemented in Monocle 3-alpha (resolution = 10−4 for most clustering analysis). (10) Subclusters that expressed low levels of the genes that were found to be differentially expressed in step 5, had high levels of markers specific to a different partition, and had relatively high doublet scores, were labelled as doublet-derived subclusters and removed from the analysis. On average, this procedure eliminated 3.4% of cells from each experiment (range 0.5–13.2%) of the cells in each experiment (Extended Data Fig. 2a–e).

    Cell clustering and cell-type annotations

    For data from individual experiments, after removing the potential doublets detected by the above three steps, we further filtered out the potential low-quality cells by investigating the numbers of UMIs and the proportion of reads mapping to the exonic regions per cell (Extended Data Fig. 2f). Then, we merged cells from individual experiments to generate the penultimate dataset, which included 15 sci-RNA-seq3 experiments and 21 runs of the Illumina NovaSeq instrument. In our early embeddings of this penultimate dataset, we noticed that one mouse embryo at E14.5 had a grossly reduced proportion of neuronal cells. This particular sample had been divided during pulverization, and we suspect that specific anatomical portions of the frozen embryo did not make it into the experiment. We therefore removed cells from this E14.5 embryo, and we further filtered out cells from the whole dataset with doublet score (by Scrublet) > 0.15 (~0.3% of the whole dataset), as well as cells with either the percentage of reads mapping to ribosomal chromosome (Ribo%) > 5 or the percentage of reads mapping to mitochondrial chromosome (Mito%) > 10 (~0.1% of the whole dataset). Finally, 11,441,407 cells from 74 embryos were retained, of which the median UMI count per cell is 2,700 and median gene count detected per cell is 1,574. For this final matrix, the number of cells recovered by each embryo and the basic quality information for cells from each sci-RNA-seq3 experiment is summarized in the Supplementary Tables 1 and 2. For sex separation and confirmation of embryos with or without sex genotyping, we counted reads mapping to a female-specific non-coding RNA (Xist) or chromosome Y genes (except Erdr1 which is in both chromosome X and chromosome Y). Embryos were readily separated into females (more reads mapping to Xist than chromosome Y genes) and males (more reads mapping to chromosome Y genes than Xist).

    We then applied Scanpy v.1.6.020 to this final dataset, performing conventional single-cell RNA-seq data processing: (1) retaining protein-coding genes, lincRNA, and pseudogenes for each cell and removing gene counts mapping to sex chromosomes; (2) normalizing the UMI counts by the total count per cell followed by log transformation; (3) selecting the 2,500 most highly variable genes and scaling the expression of each to zero mean and unit variance; (4) applying PCA and then using the top 30 principal components to calculate a neighbourhood graph (n_neighbors = 50), followed by Leiden clustering (resolution = 1); (4) performing UMAP visualization in 2D or 3D space (min.dist = 0.1). For cell clustering, we manually adjusted the resolution parameter towards modest overclustering, and then manually merged adjacent clusters if they had a limited number of DEGs relative to one another or if they both highly expressed the same literature-nominated marker genes. For each of the 26 major cell clusters identified by the global embedding, we further performed a sub-clustering with the similar strategies, except setting n_neighbors = 30 when calculating the neighbour graph and min_dist = 0.3 when performing the UMAP. Subsequently, we annotated individual cell clusters identified by the sub-clustering analysis using at least two literature-nominated marker genes per cell-type label (Supplementary Table 5).

    To be clear, we have hierarchically nominated three levels of cell-type annotations in the manuscript. (1) In the global embedding involving all 11.4 M cells we identified 26 major cell clusters (Fig. 1b,c and Supplementary Table 4). (2) For individual major cell clusters, we performed sub-clustering, resulting in 190 cell types (Extended Data Fig. 3 and Supplementary Table 5). (3) For a handful of cell types, in specific parts of the manuscript, we performed further sub-clustering, to identify cell subtypes. For example: (i) we re-embedded 745,494 cells from the lateral plate and intermediate mesoderm derivatives, identifying 22 subtypes, most of which correspond to different types of mesenchymal cells (Fig. 3d and Supplementary Table 12). (ii) we re-embedded 296,020 cells (glutamatergic neurons, GABAergic neurons, spinal cord dorsal progenitors and spinal cord ventral progenitors) from stages <E13, identifying 18 different neuron subtypes (Fig. 4e and Supplementary Table 12).

    Of note, we processed and analysed the birth series dataset (n = 962,697 nuclei after removing low-quality cells and potential doublets cells) and the early versus late somites data (n = 104,671 nuclei after removing low-quality cells and potential doublets cells) using exactly the same strategy, except without performing sub-clustering on each major cell cluster.

    Whole-mouse embryo analysis

    Each cell was assigned to the mouse embryo from which it derived based on its reverse transcription barcode. For each of the 74 samples, UMI counts mapping to the sample were aggregated to generate a pseudo-bulk RNA-seq profile for the sample. Each cell’s counts were then normalized by dividing by its estimated size factor. The data were then log2-transformed after adding a pseudocount, and PCA was performed on the transformed data using the 3,000 most highly variable genes. The normalization and dimension reduction were performed using Monocle v3.

    Quantitatively estimating cell number for individual mouse embryo at any stage during organogenesis

    To estimate the cell number of individual embryos, we selected a representative embryo from 12 timepoints at 1-day increments, from E8.5 to P0 (roughly considered as E19.5). Each embryo was digested with proteinase K overnight, and total genomic DNA was isolated with a Qiagen Puregene tissue kit (Qiagen 158063). DNA was quantified and cell number was estimated by taking the total ng of recovered DNA and assuming 2.5 billion base pairs per mouse genome (times two for a diploid cell), 650 g per mole of a base pair. Estimating cell number this way does not include any losses due to the DNA preparation, and does not count non-nucleated cells.

    Based on the experimentally estimated cell numbers of those 12 embryos, we applied polynomial regression (degree = 3) to fix a curve across embryos between the embryonic day and log2-scaled cell number (adjusted R2 > 0.98) (Extended Data Fig. 2l). P0 was treated as E19.5 in the model. Then, the total cell number of a whole mouse embryo at any day between E8.5 and P0 is predicted using the below formula:

    $${\log }_{2}({\rm{cell}}\,{\rm{number}})=0.011369\times {{\rm{day}}}^{3}-0.583861\times {{\rm{day}}}^{2}+10.397036\times {\rm{day}}-35.469755$$

    To estimate the dynamic ‘doubling time’ of the total cell number in a whole mouse embryo, at a given timepoint (day), we took the derivative from the above formula as the log2-scaled proliferation rate p(day), and then calculated \(24\times 2/{2}^{p({\rm{day}})}\), resulting in a point estimate of the number of hours required for the mouse embryo to double its total cell number (Extended Data Fig. 2m).

    Characterizing transcriptional heterogeneity in the posterior embryo

    We re-analysed 121,118 cells which were initially annotated as NMPs and spinal cord progenitors, mesodermal progenitors (Tbx6+), notochord, ciliated nodal cells, or gut, from embryos during the early somitogenesis (somite counts 0–34; E8–E10). Three clusters were identified, with cluster 1 dominated by NMPs and their derivatives (n = 98,545 cells), cluster 2 dominated by notochord and ciliated nodal cells (n = 3,949 cells), and cluster 3 dominated by gut cells (n = 18,624 cells).

    To characterize transcriptional heterogeneity within each of the three cell clusters, we performed PCA on the 2,500 most highly variable genes in each cluster. Then, we calculated the Pearson correlation between the expression of the top highly variable genes and each of the top principal components within each of the three cell clusters. In brief, for each cell cluster, the top 2,500 highly variable genes were identified and their gene expression values were calculated from original UMI counts normalized to total UMIs per cell, followed by natural-log transformation and scaling. After performing Pearson correlation with the selected principal component, significant genes were identified if their correlation coefficients are less than mean − 1 × s.d. or greater than mean + 1 × s.d. of all the correlation coefficients, and false discovery rate < 0.05. In addition, we identified differentially expressed genes between early (n = 4,949 cells) and late (n = 3,910 cells) NMPs, using the FindMarkers function of Seurat v363, after filtering out genes that are detected in <10% of cells in both of the two populations. Significant genes were identified if their absolutely log-scaled fold changes >0.25, and adjusted P values < 0.05. Of note, here cells are labelled as NMPs if they are both strongly T+ (raw count ≥5) and Meis1 (raw count = 0).

    In Fig. 2k, the Pearson correlation coefficient between gene expression for the top highly variable genes and either PC1 of notochord (x axis) or PC1 of gut (y axis) are plotted. The overlapped genes between two cell clusters are shown as each dot, and the overlapped significant genes are highlighted in blue. The first quadrant corresponds to the inferred anterior aspect of each cluster, while the third quadrant corresponds to the inferred posterior aspect. In Fig. 2l, the log-scaled fold change of the average expression for the top highly variable genes between early versus late NMPs (x axis), and the Pearson correlation coefficient between gene expression for the top highly variable genes and PC2 of gut (y axis) are plotted. The first quadrant is associated with early somite counts for each cluster, while the third quadrant is associated with late somite counts. In the gene expression line plots in Fig. 2e, left and Fig. 2k,l, right, gene expression values were calculated from original UMI counts normalized to total UMIs per cell, followed by natural-log transformation. The line of gene expression was plotted by the geom_smooth function in ggplot2.

    Spatial mapping with Tangram

    To infer the spatial origin of each lateral plate and intermediate mesoderm derivative, we used a public dataset called Mosta46, which profiles spatial transcriptomes for 53 sections of mouse embryos spanning 8 timepoints from E9.5 to E16.5. We combined this data with our own data to perform spatial mapping analysis using Tangram47. In brief, for each timepoint of the Mosta data, we combined scRNA-seq data from three adjacent timepoints from our data (for example, E16.25, E16.5 and E16.75 from scRNA-seq versus E16.5 from Mosta data), and the total number of voxels within each section was randomly downsampled to 9,000 for computational efficiency. We used the Tangram with default parameters to estimate the spatial coordinates of cells from each cell type in the scRNA-seq data, and then visualized the results on the coordinates provided by Mosta. The Tangram model was trained in GPU mode using a NVIDIA A100 GPU. After applying Tangram, for each section, a cell-by-voxel matrix with mapping probabilities was returned. This matrix shows the probability that each cell originated from each voxel in the section. To reduce noise, we further smoothed the mapping probabilities for each voxel by averaging values of their k-nearest neighbouring voxels (k is calculated by natural-log-scaled total number of voxels on that section) followed by scaling it to 0 to 1 across voxels of each section. Although only selected results are presented in the paper, the mapping results for each Mosta section on which we performed this analysis are available at https://github.com/ChengxiangQiu/JAX_code/blob/main/spatial_mapping.tar.gz.

    Generating a cell-type tree for mouse development

    We collected and combined scRNA-seq data from four published datasets, which consisted of 110,000 cells spanning E0 to E8.5, and the main dataset described in this paper, which consisted of 11.4 million cells spanning E8 to P0 (Supplementary Table 17). We generated the tree of cell types for mouse development via the following steps.

    First, based on data source, developmental window and cell-type annotations, we split cells into fourteen subsystems which could be separately analysed and subsequently integrated. The first two subsystems correspond to the pre-gastrulation and gastrulation phases of development and are based on the external datasets4,5,6,7. The remaining 12 subsystems derive from the data reported here, and collectively encompass organogenesis and fetal development (Supplementary Tables 17 and 18).

    Second, dimensionality reduction was performed separately on cells from each of the fourteen subsystems. Manual re-examination of each subsystem led to some corrections or refinements of cell-type annotations, ultimately resulting in 283 annotated cell-type nodes, some with only a handful of cells (for example, 60 ciliated nodal cells) and others with vastly more (for example, 650,000 fibroblasts) (Supplementary Tables 19 and 20). Of note, each of these annotated cell-type nodes derives from one data source, such that there are some redundant annotations that facilitate ‘bridging’ between datasets (Extended Data Fig. 11d–h). In contrast to our previous strategy in which nodes were stage-specific8, each cell-type node here is temporally asynchronous, and of course may also contain other kinds of heterogeneity (for example, spatial, differentiation, cell cycle and others).

    Third, we sought to draw edges between nodes (Fig. 5a–f). Within each subsystem, we identified pairs of cells that were MNNs in 30-dimensional PCA space (k = 10 neighbours for pre-gastrulation and gastrulation subsystems, k = 15 for organogenesis and fetal development subsystems). Although the overwhelming majority of MNNs occurred within cell-type nodes, some MNNs spanned nodes and are presumably enriched for bona fide cell-type transitions. To approach this systematically, we calculated the total number of MNNs that spanned each possible pair of cell-type nodes within a given subsystem, normalized by the total number of possible MNNs between those nodes, and ranked all possible intra-subsystem edges based on this metric (Supplementary Table 21). Of note, due to its complexity, this was done in two stages for the ‘Brain and spinal cord’ subsystem, first applying the heuristic to the subset of cell types corresponding to the patterned neuroectoderm, and then again to identify edges between the patterned neuroectoderm and its derivatives (that is, neurons, glial cells and others).

    Fourth, we manually reviewed the ranked list of 1,155 candidate edges for biological plausibility (those with a normalized MNN score > 1; Extended Data Fig. 11d), resulting in 452 edges which we manually annotated as more likely to correspond to either ‘developmental progression’ or ‘spatial continuity’ (Supplementary Table 22). Where nodes were connected to more than one other node, distinct subsets of cells were generally involved in each edge (Fig. 5a,b,d,e), and inter-node MNN pairs exhibited temporal coincidence (Fig. 5c,f). As only a handful of cells were profiled in the pre-gastrulation subsystem, those edges were added manually.

    Finally, to bridge subsystems, we performed batch correction and co-embedding of selected timepoints from either the pre-gastrulation and gastrulation datasets, or the gastrulation and organogenesis and fetal development datasets, to identify equivalent cell-type nodes, resulting in a third category of ‘dataset equivalence’ edges (Extended Data Fig. 11e–h). For example, we performed anchor-based batch correction63 followed by integration between cells from E6.5 to E8.5 generated on the 10x Genomics platform7 (n = 108,857 cells) and the earliest 1% of this dataset (0–12 somite stage embryos) generated by sci-RNA-seq3 (n = 153,597 nuclei) (Extended Data Fig. 11e,f). This allowed us to identify 36 cell types from the integrated dataset, which we used to identify bridging edges between the gastrulation subsystem and the later subsystems (Extended Data Fig. 11g,h). Most of the 12 organogenesis and fetal development subsystems originate in cell-type nodes for which equivalent nodes are already present at gastrulation. The exceptions, presumably due to undersampling of this transition, were the ‘blood’ and ‘PNS neuron’ subsystems, for which we manually added edges to connect them with biologically plausible pseudo-ancestors. Altogether, we added 55 inter-subsystem edges.

    In practice, a small number of nodes in the tree have more than one parent, so the ‘tree’ is formally a rooted, directed graph that represents mouse development from E0 to P0. The visualization shown in Fig. 5g was created using yFiles Hierarchical layout in Cytoscape v3.9.1. For presentation purposes, we removed most of the spatial continuity edges, except for those between spinal cord dorsal and ventral progenitors after E13.0 and GABAergic and glutamatergic neurons after E13.0. We also merged nodes with redundant labels derived from different datasets (that is, dataset equivalence edges). This resulted in a rooted graph with 262 cell-type nodes and 338 edges.

    Our evaluation of the robustness of our approach to technical factors or parameter choices is provided in Extended Data Fig. 11a–c and Supplementary Note 2.

    Nominating key transcription factors and genes

    The list of 1,636 mouse proteins that are putatively transcription factors was collated from AnimalTFDB v3 (http://bioinfo.life.hust.edu.cn/AnimalTFDB/)78. For each edge in the cell-type tree, we stratified each cell-type transition into four phases. Specifically, we identified the subset of cells within each node that were either ‘inter-node’ MNNs of the other cell-type or ‘intra-node’ MNNs of those cells. If A → B, this approach effectively models the transition as group 1 → 2 → 3 → 4 (Extended Data Fig. 11i,j). Next, we identified DETFs and genes (DEGs) across each portion of the modelled transition—that is, early (1 → 2), inter-node (2 → 3) and late (3 → 4)—by applying FindMarkers function in Seurat v3 with parameters (logfc.threshold = 0, min.pct = 0). This strategy highlights differences between cells that are most proximate to the cell-type transition itself.

    After excluding dataset equivalence edges and the ‘pre-gastrulation’ subsystem, we nominated key transcription factors and genes that specify cell types for each of the 436 edges. Of note, the directionality of many of these edges was not immediately obvious (that is, those annotated as “spatial continuity” edges). In these cases, the orientation of the ‘early’ and ‘late’ phases is arbitrary. For edges with a relatively small number of MNN pairs, we expanded each group to at least 200 cells by iteratively including their MNNs within the same cell type, to increase statistical power.

    Identifying cell types with abrupt transcriptional changes before versus after birth

    To systematically identify which cell types exhibit abrupt transcriptional changes before versus after birth, we performed the following steps.

    • We focused on the 71 cell types with at least 200 cells from P0 and at least 200 cells from at least 5 timepoints prior to P0.

    • We combined cells from animals collected subsequent to E16 and performed PCA based on the top 2,500 highly variable genes.

    • Timepoints with at least 200 cells were selected and cells were downsampled from each timepoint to the median number of cells across those selected timepoints.

    • The k-nearest neighbours (k was adjusted for different cell types, by taking the log2-scaled median number of cells across the selected timepoints) were identified in PCA space (n = 30 dimensions).

    • We calculated the average proportion of nearest neighbour cells that were from a different timepoint for cells within each cell type. In this framing, a low proportion of neighbours from different timepoints corresponds to a relatively abrupt change in transcriptional state.

    We subjected the birth-series dataset to a similar analysis. For each major cell cluster in the birth-series dataset, we took cells from the 6 pups delivered by C-section and calculated the Pearson correlation coefficient between the timepoint of each cell and the average timepoints of its 10 nearest neighbours identified from the global PCA embedding (n = 30 dimensions). In this framing, a high correlation indicates that the cell and its nearest neighbours all underwent rapid, synchronized changes in transcriptional state.

    Reporting summary

    Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

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