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LAP2alpha maintains a mobile and low assembly state of A-type lamins in the nuclear interior

  1. Nana Naetar  Is a corresponding author
  2. Konstantina Georgiou
  3. Christian Knapp
  4. Irena Bronshtein
  5. Elisabeth Zier
  6. Petra Fichtinger
  7. Thomas Dechat
  8. Yuval Garini
  9. Roland Foisner  Is a corresponding author
  1. Max Perutz Labs, Center for Medical Biochemistry, Medical University of Vienna, Vienna Biocenter Campus (VBC), Austria
  2. Physics Department and Nanotechnology Institute, Bar Ilan University, Israel
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Cite this article as: eLife 2021;10:e63476 doi: 10.7554/eLife.63476

Abstract

Lamins form stable filaments at the nuclear periphery in metazoans. Unlike B-type lamins, lamins A and C localize also in the nuclear interior, where they interact with lamin-associated polypeptide 2 alpha (LAP2α). Using antibody labeling, we previously observed a depletion of nucleoplasmic A-type lamins in mouse cells lacking LAP2α. Here, we show that loss of LAP2α actually causes formation of larger, biochemically stable lamin A/C structures in the nuclear interior that are inaccessible to lamin A/C antibodies. While nucleoplasmic lamin A forms from newly expressed pre-lamin A during processing and from soluble mitotic lamins in a LAP2α-independent manner, binding of LAP2α to lamin A/C during interphase inhibits formation of higher order structures, keeping nucleoplasmic lamin A/C in a mobile state independent of lamin A/C S22 phosphorylation. We propose that LAP2α is essential to maintain a mobile lamin A/C pool in the nuclear interior, which is required for proper nuclear functions.

Introduction

Lamins are intermediate filament proteins in metazoan nuclei that, together with numerous inner nuclear membrane proteins, form a filamentous protein meshwork at the nuclear periphery, called the nuclear lamina (Gruenbaum and Foisner, 2015a). Based on their biochemical properties and expression patterns, lamins are grouped into two subtypes, A-type and B-type lamins. B-type lamins are ubiquitously expressed in all cell types and throughout development (Yang et al., 2011), whereas A-type lamins are expressed at low levels in embryonic stem cells and undifferentiated cells, but are significantly upregulated during differentiation (Constantinescu et al., 2006; Eckersley-Maslin et al., 2013; Röber et al., 1989). In mammalian cells, the major A-type lamins are lamins A and C encoded by the Lmna gene, whereas the two major B-type lamins, lamins B1 and B2, are encoded by Lmnb1 and Lmnb2, respectively (Gruenbaum and Foisner, 2015a). Both lamin subtypes share a similar intermediate filament protein-type domain structure with a central rod domain, an N-terminal head and a globular C-terminal tail, which contains a nuclear localization signal, an Ig fold and, except for lamin C, a C-terminal CaaX motif (C: cysteine; a: aliphatic amino acid; X: any amino acid) (de Leeuw et al., 2018; Gruenbaum and Medalia, 2015b). The CaaX motif undergoes a series of post-translational modifications, including farnesylation of the cysteine, removal of the last three amino acids, followed by carboxymethylation (Rusiñol and Sinensky, 2006). Whereas B-type lamins remain farnesylated and carboxymethylated, pre-lamin A undergoes an additional processing step catalyzed by the metalloprotease Zmpste24, leading to the removal of 15 amino acids from its C-terminus, including the farnesylated cysteine residue. As a consequence, mature B-type lamins are tightly associated with the nuclear membrane via their farnesylated C-terminus and mainly localize at the nuclear periphery, whereas mature lamin A is found both in filaments within the peripheral nuclear lamina, and additionally in a soluble and dynamic pool in the nuclear interior (Naetar et al., 2017). Lamin C that lacks a CaaX motif contributes also to the peripheral and nucleoplasmic pool of A-type lamins.

Lamin filaments at the nuclear periphery were recently visualized by cryo-electron tomography (Turgay et al., 2017), but their structure and assembly are far from being well understood. Lamins form dimers via their central rod domains, which further assemble into head-to-tail polymers (de Leeuw et al., 2018). Recent work by the Medalia lab using cryo-electron tomography revealed that in mammalian nuclei two head-to-tail filaments assemble laterally into 3.5-nm-thick filaments in a staggered fashion (Turgay et al., 2017). Lamin filaments at the nuclear periphery are considered stable, resistant to biochemical extraction and highly immobile (Bronshtein et al., 2015; Moir et al., 2000; Shimi et al., 2008). Together with proteins of the inner nuclear membrane, they fulfill essential functions, defining the mechanical properties of nuclei (Buxboim et al., 2014; Cho et al., 2019; Davidson and Lammerding, 2014; Osmanagic-Myers et al., 2015; Swift et al., 2013) and regulating higher order chromatin organization through anchorage of peripheral heterochromatic genomic regions (Solovei et al., 2013; van Steensel and Belmont, 2017).

The regulation and properties of A-type lamins in the nuclear interior are far less understood, although recent studies have unraveled novel functions of lamins in the nuclear interior in chromatin regulation that are fundamentally different from those fulfilled by the nuclear lamina (Gesson et al., 2014; Naetar et al., 2017). Nucleoplasmic lamins A and C bind to and regulate euchromatic regions of the genome, globally affecting epigenetic modifications and possibly chromatin accessibility (Gesson et al., 2016; Naetar et al., 2017). They also provide chromatin ‘docking sites’ in the nucleoplasm slowing down chromatin movement in nuclear space (Bronshtein et al., 2015). Moreover, intranuclear A-type lamins are required for the proper assembly of repressive polycomb protein foci (Bianchi et al., 2020; Cesarini et al., 2015) and are involved in the regulation of telomere function (Chojnowski et al., 2015; Gonzalez-Suarez et al., 2009; Wood et al., 2014). Nucleoplasmic lamins were also found to affect gene expression directly by binding to gene regulatory sequences, such as promoters and enhancers (Briand et al., 2018; Ikegami et al., 2020; Oldenburg et al., 2017).

The structure and assembly state of nucleoplasmic lamins remain enigmatic. Fluorescence recovery after photobleaching (FRAP), continuous photobleaching (CP), and fluorescence correlation spectroscopy (FCS) studies of fluorescently tagged A-type lamins showed that the majority of nucleoplasmic lamin complexes are highly mobile compared to the stable peripheral lamina (Bronshtein et al., 2015; Moir et al., 2000; Shimi et al., 2008). Biochemical studies revealed that intranuclear lamins can be easily extracted in salt and detergent-containing buffers, suggesting that they have a low assembly state (Kolb et al., 2011; Naetar et al., 2008) and/or weaker interactions with nuclear protein complexes and chromatin compared to the lamins at the lamina. Taken together, the dynamic nucleoplasmic lamin pool has very distinct properties compared to the static and stable peripheral lamina, allowing them to fulfill a unique set of functions. However, why and how nucleoplasmic A-type lamins display such different properties in the nuclear interior compared to their counterparts at the nuclear lamina remains poorly understood. Possible mechanisms include post-translational modifications and/or interactions with specific nucleoplasmic binding partners. For example, phosphorylation of lamins influences the localization, solubility and mobility of lamins both during mitosis and interphase (Kochin et al., 2014; Machowska et al., 2015). Moreover, the specific interaction partner of nucleoplasmic lamins, lamin-associated polypeptide 2 alpha (LAP2α) was suggested to regulate the localization of lamin A/C in the nuclear interior, since the nucleoplasmic lamin pool was found significantly reduced in the absence of LAP2α (Dechat et al., 2000; Gesson et al., 2016; Gesson et al., 2014; Naetar et al., 2008). However, the mechanisms remain unknown.

Here, we address the open question if and how LAP2α regulates the formation, properties and functions of nucleoplasmic A-type lamins. Surprisingly, we find that LAP2α is not involved in the generation of the nucleoplasmic lamin A/C pool from newly synthesized lamin A/C in interphase cells and from mitotic lamins in G1 phase, but instead regulates lamin A/C properties in a lamin A/C phosphorylation-independent manner. These findings led to a new perspective of LAP2α function in lamin A/C regulation, suggesting that LAP2α affects the state of lamin A/C assembly and/or their interactions in the nucleoplasm. We show that in the absence of LAP2α, lamins A and C in the nuclear interior are more resistant toward biochemical extraction, possibly through the formation of higher order structures and/or stronger interactions with nuclear components, leading to an impaired detection by antibodies and decreased mobility. The impaired accessibility for antibodies against the lamin A/C N-terminus likely explains the previously observed reduction of nucleoplasmic lamins A/C in cells and tissues lacking LAP2α (Naetar et al., 2008). We propose that the interaction of soluble lamins with LAP2α in the nuclear interior maintains their mobile, low assembly state, which is important for proper lamin functions in nuclear organization and chromatin regulation.

Results

The nucleoplasmic lamin pool forms independently of LAP2α through at least two different processes

Previous observations using antibody labeling revealed a strong reduction of nucleoplasmic A-type lamins in cells lacking LAP2α, suggesting a prominent role for LAP2α in the formation or regulation of intranuclear lamins (Naetar et al., 2008). In order to elucidate the potential role of LAP2α in the formation of the nucleoplasmic pool of A-type lamins in interphase cells and during nuclear reassembly after mitosis, we performed live cell imaging of wildtype and LAP2α knockout HeLa cells (generated by CRISPR-Cas9, see Figure 1—figure supplement 1). First, we expressed GFP-pre-lamin A and imaged emerging pre-lamin A structures 5 hr post-transfection, when the newly expressed pre-lamin A undergoes post-translational processing (Figure 1A, columns 1 and 2, see also video files 1–3 associated with Figure 1). The nascent, transiently farnesylated pre-lamin A initially localizes to the nuclear periphery with very little visible nucleoplasmic staining (Figures 1A, 0-20’). Time-resolved quantification of the nucleoplasmic over peripheral fluorescence signal (Figure 1A, lower panel) reveals that lamins in the nuclear interior emerge gradually within 20 min, probably after their release from the nuclear periphery during further processing and removal of the C-terminal farnesyl group. Accordingly, an ectopically expressed, farnesylation-deficient mature lamin A (lacking its C-terminal 15 amino acids) is detectable prominently in the nucleoplasm at early timepoints when lamin structures first emerge (Figure 1—figure supplement 2A, see also video file 9 associated with Figure 1—figure supplement 2). Furthermore, an assembly-deficient lamin AΔK32 mutant (Bank et al., 2011; Bertrand et al., 2012) expressed in its pre-lamin form appears first at the nuclear periphery, before it translocates completely into the nucleoplasm (Figure 1A, third panel, see also video file 4 associated with Figure 1), while the farnesylation-deficient mature form of lamin AΔK32 mutant is exclusively in the nuclear interior throughout imaging (Figure 1—figure supplement 2A, see also video file 10 associated with Figure 1—figure supplement 2). These data suggest that post-translational processing rather than lamin filament assembly is the driving force for the initial localization of nascent pre-lamin A to the nuclear periphery.

Figure 1 with 14 supplements see all
Absence of LAP2α does not affect formation of a nucleoplasmic pool of exogenously expressed GFP-lamin A.

(A) Wildtype (WT) and LAP2α knockout (KO) HeLa cells (see Figure 1—figure supplement 1) were transiently transfected with EGFP-pre-lamin A or EGFP-ΔK32 pre-lamin A as indicated and analyzed by live-cell imaging 5 hr post-transfection. Schematic drawing on top explains lamin A processing state at the time of imaging. Time is indicated in minutes. See also video files: Figure 1—videos 1 and 2, corresponding to panels 1 and 2, Figure 1—video 3 displaying an identically treated second HeLa LAP2α knockout clone, and Figure 1—video 4, corresponding to panel 3. Bottom: Ratio of nucleoplasmic to peripheral mean signal intensity of EGFP-lamin A was calculated (see Materials and methods for details) and normalized to time point ‘T’. ‘T’ defines the moment preceding a significant increase of nucleoplasmic lamin A in each cell. ‘T+/-x’ indicates imaging time relative to ‘T’ with ‘x’ corresponding to minutes. Dashed lines indicate measurements performed in a smaller number of cells (nwt = 6, nko = 6), demonstrating that nucleoplasmic lamin levels remain steadily low prior to the initial release of lamin A into the nucleoplasm (time point ‘T’). Graphs display mean values ± S.E.M (nwt = 18, nko = 23). p Values (repeated measures two-way ANOVA) are ptime <0.0001 and pgenotype = 0.2160 (non-significant). (B) WT and LAP2α KO HeLa cells were transfected as in (A), but imaged 24 hr post-transfection, where ectopically expressed lamin A is largely matured. See also video files: Figure 1—videos 5 and 6, corresponding to panels 1 and 2, Figure 1—video 7 displaying an identically treated second HeLa LAP2α knockout clone, and Figure 1—video 8, corresponding to panel 3. Graphs on the right present examples for GFP-lamin A fluorescence intensity measured along the dashed lines in the images (20’ and 240’). Bottom: Nucleoplasmic over peripheral signal ratio was determined using line profiles for all time points (see Materials and methods for details), normalized to mitosis/early G1 (time point 0’) and plotted versus imaging time. Graphs display mean values ± S.E.M. (nwt = 10, nko = 12). p Values (repeated measures two-way ANOVA) ptime <0.0001 and pgenotype = 0.2564 (non-significant). Scale bar: 10 µm.

In order to test whether LAP2α is involved in the subsequent release of the newly synthesized, processed lamin A from the lamina into the nuclear interior, we imaged nascent GFP-pre-lamin A in LAP2α knockout cells (Figure 1A). The increase in the lamin A signal in the nuclear interior occurred with similar kinetics in LAP2α knockout and wildtype cells, suggesting that fully processed lamin A/C can translocate to the nuclear interior independent of LAP2α.

To test whether LAP2α may affect the dynamic behavior of mature lamin A during the cell cycle, we analyzed GFP-pre-lamin A-expressing wildtype and LAP2α knockout HeLa cells 24 hr post transfection, when the majority of the ectopic pre-lamin A has been processed into the mature form (Figure 1B, see also video files 5–7 associated with Figure 1). As previously reported (Broers et al., 1999; Moir et al., 2000), lamin A structures completely disassemble at the onset of mitosis (Figure 1B, 0’), and, during nuclear reassembly, lamin A localizes uniformly throughout the nucleus in late telophase / early G1 phase (Figure 1B, 20’). Lamin A levels in the nuclear interior then gradually decrease within around 200 min due to relocation of the majority of lamins to the nuclear periphery, but a fraction of lamin A remains in the nuclear interior throughout interphase (Figure 1B, 40-240’, see also Dechat et al., 2000; Naetar et al., 2008). As expected, ectopically expressed mature GFP-lamin A shows the same cell cycle-dependent behavior (Figure 1—figure supplement 2B, see also video file 11 associated with Figure 1—figure supplement 2). However, an assembly-deficient mutant, lamin AΔK32 remains fully nucleoplasmic throughout G1 phase, independent of whether the pre- or mature form was expressed (Figure 1B and Figure 1—figure supplement 2B, see also video files 8 and 12). Thus, lamin filament assembly is essential for the accumulation of A-type lamins at the nuclear periphery during G1 phase. In order to test the effect of LAP2α on the redistribution of lamins from the nuclear interior to the periphery after mitosis, we performed live cell imaging of LAP2α knockout cells. Quantification of the nucleoplasmic over peripheral GFP-lamin A signal after mitosis revealed a similar gradual decrease in nucleoplasmic staining during G1 phase in LAP2α knockout and wildtype cells, and more surprisingly, the steady state levels of lamin A in the nuclear interior throughout interphase were unaffected in LAP2α knockout versus wildtype cells (Figure 1B, lower panel).

In conclusion, we show that nucleoplasmic A-type lamins are generated by two different pathways: (i) newly synthesized pre-lamin A redistributes from the periphery to the nuclear interior upon post-translational processing and (ii) a fraction of soluble A-type lamins of the previous mitosis remains in the nucleoplasm during lamina assembly in G1 phase. However, none of these processes were significantly altered in the absence of LAP2α, suggesting that nucleoplasmic A-type lamins form independently of LAP2α. In addition, steady-state lamin A levels in the nuclear interior in interphase apparently were unchanged in the absence of LAP2α. These findings are conflicting with our previous results showing loss of nucleoplasmic lamin A/C in LAP2α knockout cells in immunofluorescence microscopy (Naetar et al., 2008).

LAP2α-deficiency does neither affect the formation nor the steady state level of endogenous lamin A in the nuclear interior

We reasoned that the lack of any effect of LAP2α loss on the formation of the nucleoplasmic lamin A pool may be due to overexpression of the ectopic GFP-lamin A. Thus, in order to create a more physiological setting, we tagged the endogenous lamin A gene with mEos3.2, a monomeric variant of the photoconverter EosFP (Zhang et al., 2012), in wildtype and LAP2α knockout mouse fibroblasts using CRISPR-Cas9 (Figure 2A, Figure 2—figure supplements 1 and 2). Following a rigid testing of the wildtype clones (see details in Figure 2—figure supplement 1), we picked a clone (WT clone #21) for further analyses, which is tetraploid and expresses tagged lamin A/C from 1 allele and untagged lamin A/C from three alleles in a 1:3 ratio at the transcript level (Figure 2B,C and Figure 2—figure supplement 1F,G). At the protein level, this ratio shifts to 1:8, suggesting that the addition of the mEos3.2-tag leads to a slight destabilization of lamin A and C protein (Figure 2—figure supplement 1G). However, the tagged lamins behave like untagged proteins (see below) and the level of tagged lamin A/C proteins was sufficient for high-quality microscopic analysis (Figure 2C–H). We then generated isogenic LAP2α knockout clones from this WT clone by CRISPR-Cas9 (for details see Figure 2—figure supplement 2). Both WT and LAP2α knockout clones display normal nuclear morphology and the mEos3.2-tagged lamin A/C correctly localizes at the nuclear periphery and in the nuclear interior (Figure 2C) and was stably integrated into the nuclear lamina, as demonstrated by photoconversion of tagged lamins, where no significant mobility of photo-converted lamins in the lamina was observed for several hours (Figure 2D). Moreover, the solubility in high-salt and detergent-containing buffers was comparable between untagged and tagged lamins A and C (Figure 2—figure supplement 1G). Thus, endogenously tagged lamin A/C behaves like untagged protein in terms of localization, dynamics and biochemical properties (Figure 2 and Figure 2—figure supplement 1), supporting its functionality. Notably, cells with all Lmna alleles tagged, thus expressing only mEos3-2-lamin A/C, displayed similar viability, nuclear morphology, and lamina structure when compared to parental control cells expressing only WT lamin A/C (Figure 2—figure supplement 1H and I).

Figure 2 with 3 supplements see all
Absence of LAP2α does not alter endogenous nucleoplasmic lamin A/C levels.

(A) Schematic view of exons 1 and 2 (bars) and adjacent introns of the mouse Lmna locus (top), the targeting construct provided as homology-directed repair template after Cas9-mediated double strand break (middle), and the knock-in allele after successful integration of mEos3.2 (bottom). Very long introns are not displayed in their original length as indicated by a double slash. The second, light-colored exon 1 encodes the N-terminus of meiosis-specific lamin C2. The target sequence of Lmna-specific sgRNA1 in exon 1 is shown (blue). Protospacer-adjacent motif (PAM) is marked in red. Red arrowhead: expected Cas9 cut site. HA-L: Homology Arm-Left; HA-R: Homology Arm-Right. For further cell characterization see Figure 2—figure supplement 1. (B) mEos3.2-Lmna clone derived from wildtype (WT) mouse dermal fibroblasts (MDFs) and subclones generated after treatment with LAP2α-specific sgRNAs 1 or 3 (see Figure 2—figure supplement 2) were processed for western blotting using antibodies to the indicated antigens (anti lamin A/C E1, anti LAP2α 1H11). Positions of mEos-tagged and untagged lamins A and C are indicated. Wildtype and LAP2α knockout (KO) MDFs were added as controls. (C) mEos3.2-Lmna WT and LAP2α KO cells were processed for immunofluorescence microscopy using antibodies against LAP2α (1H11) and DAPI to visualize DNA. Confocal images are shown. Bar: 10 μm. (D) mEos3.2-Lmna WT cells were exposed to UV light (405 nm) to convert mEos3.2-lamin A/C from green to red. Cells were then tracked for 14 hr by live-cell imaging to test for stable integration of mEos3.2-lamin A/C into the peripheral lamina. No significant mobility of converted lamins in the lamina was observed over the entire imaging time. Pictures display merged Z-stacks to visualize most of the peripheral lamina. Bar: 10 μm. (E) Representative example of a LAP2α KO cell expressing mEos3.2-lamin A/C tracked by live-cell imaging throughout G1, followed by automated calculation of nucleoplasmic over peripheral lamin A/C ratio (t0: first available image after mitosis). The area corresponding to the nuclear lamina is delineated by red lines and was defined by a custom-made algorithm implemented in FIJI/ImageJ software. Average values for isogenic mEos3.2-Lmna LAP2α WT and KO clones were plotted in curves (F). The designation ‘sg1’ and ‘sg3’ refers to the sgRNA used to generate that clone (see Figure 2—figure supplement 2). Clones treated with sgRNAs, but still expressing LAP2α, were included as additional controls and are designated as ‘WT sg ctrl’. The 20th and 80th percentile are displayed for each curve (shaded area). nWT = 129, nWTsg1ctrl=159, nKOsg1 = 106, nWTsg3ctrl=111, nKOsg3 = 147 (G) Isogenic clones were fixed and DNA was stained with DAPI. Nucleoplasmic (N) to peripheral (P) signal intensity of mEos3.2-Lamin A/C was calculated for single cells (green dots) using a custom-made FIJI plugin and plotted versus DAPI fluorescence intensity, indicative of cell cycle stage (see also Figure 2—source data 1). Black line outlines number of cells versus DAPI intensity (histogram). Vertical dotted lines indicate what was classified as G1 and S-G2 in (H). (H) Single-cell N/P ratios from (G) were averaged for G1 and S/G2 and are shown in a box plot (median in red within the first and third quartiles; whiskers: minimal and maximal datapoint excluding outliers). The specificity of nucleoplasmic lamin A/C signal was demonstrated by comparison to lamin B1 N/P ratios (see Figure 2—figure supplement 3).

Figure 2—source data 1

Nucleoplasmic to peripheral signal intensity ratio and DAPI fluorescence intensity of mEos3.2-Lamin A/C WT and LAP2α KO cells.

https://cdn.elifesciences.org/articles/63476/elife-63476-fig2-data1-v2.xlsx

To investigate the potential role of LAP2α in the retention of endogenous lamins A and C in the nucleoplasm in G1 phase, and in controlling steady-state levels of lamin A/C in the nuclear interior throughout interphase, we followed mEos3.2-tagged lamin A/C in wildtype and LAP2α knockout clones by live cell imaging. We performed time lapse studies starting at the exit from mitosis through G1 phase, and determined the nucleoplasmic to peripheral lamin A/C signal ratio in hundreds of cells by automated quantification using a custom-made FIJI plugin (Figure 2E). Neither the post-mitotic dynamics of nucleoplasmic lamin re-localization nor its steady-state levels in late G1 phase were affected in LAP2α knockout versus wildtype cells (Figure 2F).

To further analyze nucleoplasmic lamin A/C levels through interphase, we fixed mEos3.2-lamin A/C expressing wildtype and LAP2α knockout cells and automatically quantified their nucleoplasmic (N) to peripheral (P) lamin A/C ratio as described before. The lamin A/C N/P ratio was then plotted against the intensity of the DAPI signal to stage cells within the cell cycle (Roukos et al., 2015; Figure 2G). As expected, the nucleoplasmic to peripheral lamin A/C ratio was higher in G1 phase (WT G1:0.26 ± 0.0055) than in S/G2 phase (WT S/G2: 0.19 ± 0.0045, Dunn’s post test p<0.0001), but both ratios were unchanged in the absence of LAP2α (LAP2α KO G1: 0.29 ± 0.0054, S/G2: 0.21 ± 0.0044). The observed subtle variability is likely due to clonal differences, as it was also observed between different wildtype clones (e.g. LAP2α WT sg1 ctrl G1: 0.27 ± 0.0045, S/G2: 0.20 ± 0.0036) (Figure 2G and H). To ensure specificity of the nucleoplasmic signal and investigate the influence of out-of-focus light from the much stronger peripheral lamin A/C fluorescence, we determined N/P ratios also for lamin B1, which is exclusively localized at the periphery (Figure 2—figure supplement 3). When lamin B1 ratios, which would represent out-of-focus signal in the nucleoplasm, were substracted from lamin A/C N/P ratios, the resulting values were still significantly above 0 (Figure 2—figure supplement 3, lower panel), indicating that the detected nucleoplasmic lamin A/C signal stems, at least to a large extent, from lamins in the nuclear interior and not from out-of-focus light from the nuclear lamina.

Thus, endogenous lamins A and C are retained in the nucleoplasm after mitosis and remain in the nuclear interior throughout interphase independent of LAP2α, which is in stark contrast to previous findings demonstrating a clear reduction of nucleoplasmic lamin A/C signal in LAP2α knockout cells and tissues by immunofluorescence microscopy (Naetar et al., 2008).

LAP2α loss changes properties of nucleoplasmic lamin A/C rendering them less mobile and more resistant to biochemical extraction

In order to resolve these contradicting results, we first tested mEos3.2-lamin A/C-expressing wild-type cells and isogenic LAP2α knockout clones by immunofluorescence microscopy using an antibody to the lamin A/C N-terminus (N18), which, unlike an antibody to the lamin A/C C-terminus (3A6, see Figure 3—figure supplement 1B), preferentially stains nucleoplasmic lamins A and C (Gesson et al., 2016; Figure 3A and B, Figure 3—figure supplement 1A). While the intranuclear mEos3.2 fluorescence signal was similar in LAP2α knockout and wildtype cells, the nucleoplasmic lamin A/C antibody-derived signal was clearly reduced in LAP2α knockout versus wildtype cells (Figure 3A,B and Figure 3—figure supplement 1A, compare red and green channels and see quantification in Figure 3D). Interestingly, the intranuclear antibody staining in LAP2α knockout cells completely recovered following post-fixation treatment with DNase and RNase (Figure 3C and D), suggesting epitope masking specifically in LAP2α knockout cells.

Figure 3 with 1 supplement see all
Absence of LAP2α reduces nucleoplasmic lamin A/C staining in immunofluorescence.

(A) mEos3.2-Lmna WT and LAP2α KO sg1 cells were processed for immunofluorescence microscopy using antibody N18 against lamin A/C N-terminus, and DAPI to visualize DNA. Lamin A/C antibody N18 preferably stains nucleoplasmic lamin A/C. (B) mEos3.2-Lmna WT and LAP2α KO sg1 cells were co-cultured and processed for immunofluorescence microscopy as in (A) using antibodies N18 against lamin A/C and 1H11 against LAP2α, and DAPI to visualize DNA. Fluorescence intensity was determined for each cell along the dashed line in the red and green channel and is depicted in percent of maximum value in the graphs below. (C) LAP2α KO sg1 cells were processed as in (A) using lamin A/C antibody N18 without or with prior treatment with DNase I and RNase A to reverse N18 epitope masking. Bar: 10 μm. (D) Average nucleoplasmic fluorescence intensity from 3 wildtype and 3 LAP2α KO clones with and without prior DNase/RNase (D/R) treatment for N18 stainings and for mEos3.2 lamin A/C signal is depicted. Data represent averages ± S.E.M. Single data points are shown for each group. One-way ANOVA p value for N18 is 0.0020, F = 20.85; **p<0.01 (Tukey’s post-hoc test: WT vs. KO: p=0.0034; KO vs KO plus DNase/RNase: p=0.0033); p value for mEos3.2-lamin A/C is 0.3694 (non-significant), F = 1.181.

In order to exclude the possibility that mEos3.2-lamin A/C protein behaves differently as compared to untagged lamin A/C (both are expressed in the WT cell clone), we tested other cell clones expressing exclusively mEos3.2-tagged lamin A/C (without any untagged lamin A/C, Figure 3—figure supplement 1C and D). These clones exhibit the same properties and fitness as parental cells expressing only untagged WT lamin A/C (see Figure 2—figure supplement 1H and I) and display a similar reduction in N18 antibody-derived nucleoplasmic lamin staining in the absence of LAP2α, while the mEos3.2-lamin A/C fluorescence signal in the nuclear interior was unchanged (Figure 3—figure supplement 1C). Moreover, peripheral lamin A/C staining using the 3A6 antibody was unchanged in the absence of LAP2α in both, cells expressing only mEos3.2-lamin A/C and parental cells (Figure 3—figure supplement 1D).

Overall, these data suggest that – instead of a complete absence of nucleoplasmic A-type lamins – the reduced staining in LAP2α knockout cells reflects an alteration of nucleoplasmic lamin A/C properties that led to masking of the N-terminal epitope recognized by the antibody.

To test potential changes in nucleoplasmic lamin A/C properties in LAP2α knockout versus wildtype cells, we first tested lamin mobility. While lamins at the periphery form stable filaments, nucleoplasmic lamin A/C was shown to be highly mobile (Broers et al., 1999; Kolb et al., 2011; Shimi et al., 2008). We tested the mobility of nucleoplasmic lamins by Continuous Photobleaching (CP), where the fluorescence intensity is measured in a small spot within the nucleoplasm of mEos3.2-lamin A/C expressing cells at a high frequency over time. This analysis revealed two fluorescent sub-populations with different mobility: an immobile fast bleaching fraction, and a highly mobile slow bleaching fraction (Figure 4A, left panel). As previously demonstrated (Bronshtein et al., 2015), 10–40% of lamin A in the nucleoplasm is immobile in the majority of wildtype cells (Figure 4A). Intriguingly, in the absence of LAP2α we observed a noticeable shift toward a higher fraction of immobile lamin A/C in the nuclear interior in the range of 30 up to 60% (Figure 4A, right panel). Thus, the mobile fraction of lamin A in the nuclear interior is reduced in LAP2α knockout versus wildtype cells.

Figure 4 with 1 supplement see all
Nucleoplasmic lamin A/C is less mobile, more extraction-resistant and form larger assemblies in the absence of LAP2α.

(A) Left: Depicted is a representative curve obtained by continuous photobleaching (CP, see schematic inset on the lower left) using mEos3.2-lamin A/C WT cells, allowing to determine mobile and immobile fractions (shown on the left side of the graph) of the measured mEos3.2-lamin A/C. Blue line represents normalized average intensity of mEos3.2-lamin A/C measured over time from a selected spot inside the nucleus. Right: Immobile fractions of mEos3.2-lamin A and C protein as calculated from measured CP curves of mEos3.2-lamin A/C WT and LAP2α KO sg1 cells are depicted as histogram (see also Figure 4—source data 1). nWT = 59, nKO = 60, p=0.0053 (two-tailed student’s t test of arcsin transformed values). (B) Graph displays the diffusion co-efficient of fast and slow moving mEos3.2-lamins A and C as determined by fluorescence correlation spectroscopy (FCS) measurements (see also Figure 4—source data 2). Data represent averages ± S.E.M.; nWT (fast fraction)=106, nWT (slow fraction)=96, nKO (fast fraction)=97, nKO (slow fraction)=99; ***p<0.0001 (Mann-Whitney U test). (C) Cells from isogenic WT and LAP2α KO clones were extracted in salt and detergent-containing buffer (500 mM NaCl, 0.5% NP-40). Extracts were processed for Western blot analysis. Upper panel shows representative western blots from WT and LAP2α KO sg1 cells using antibodies against total lamin A/C (E1), lamin A/C phosphorylated at serine 22 (pS22) or LAP2α (1H11). T: total lysate; P: insoluble pellet fraction; SN: soluble, extracted supernatant fraction. Western blots were quantified and lamin signal in the supernatant was normalized to total lamin A/C. Graphs display lamins A and C levels in the supernatant fraction of LAP2α knockout samples as average fold difference ± S.E.M over wildtype samples. Single data points are shown for each group. nWT = 6, nKO = 6; **p=0.00013, ***p=2.09E-5, n.s.: non-significant (p=0.072) (paired student’s t-test on log transformed values). (D) mEos3.2-lamin A/C WT and LAP2α KO sg1 cells were processed for immunofluorescence microscopy with and without prior extraction in salt and detergent-containing buffer. Confocal super-resolution (Airyscan) images of mEos3.2-lamin A/C signal are depicted. Larger lamin A/C nucleoplasmic structures in LAP2α knockout cells are marked by arrowheads and displayed as larger inset (bottom). (E) Graphs show quantification of number (No.) of intranuclear lamin A/C structures per cell and mean area per structure. Data represent averages ± S.E.M. Left graph: nWT = 5, nKO = 7 (single data points are displayed), p=0.0076 (Mann-Whitney U test); right graph: nWT = 16, nKO = 64, p=0.0433 (Mann-Whitney U test); *p<0.05, **p<0.01.

Figure 4—source data 1

Immobile mEos3.2-lamin A/C fraction in the nucleoplasm of WT and LAP2α KO cells determined by constant photobleaching.

https://cdn.elifesciences.org/articles/63476/elife-63476-fig4-data1-v2.xlsx
Figure 4—source data 2

Diffusion co-efficient of fast and slow moving mEos3.2-lamins A and C in WT and LAP2α KO cells as determined by FCS.

https://cdn.elifesciences.org/articles/63476/elife-63476-fig4-data2-v2.xlsx

In addition, we measured the mobile lamin A/C fraction in the nuclear interior by fluorescence correlation spectroscopy (FCS), which revealed a slower and a faster-moving fraction as previously reported (Shimi et al., 2008). Interestingly, although continuous photobleaching has clearly revealed an overall decrease in the relative amount of the mobile (moving) lamin A/C pool in LAP2α knockout cells, the small remaining mobile fractions showed increased diffusion (Shimi et al., 2008). The ratio of the FCS diffusion rates (DWT/DKO, 0.76 for the fast fraction) equals the cube root of the inverted molecular weight ratio (MKO/MWT) of the nucleoplasmic lamin complex (see Materials and methods for details). Assuming that the WT complex contains a 1:1 ratio (our unpublished data) of mEos3.2-lamin A and LAP2α (100 kDa + 75 kDa), whereas LAP2α KO cells contain only mEos3.2-lamin A (100 kDa), the calculated cube root of the KO/WT molecular weight ratio is 0.83. Therefore, the measured FCS diffusions may be explained by the loss of LAP2α in the lamin A/C complex in LAP2α knockout cells (Figure 4B).

To reveal further changes in lamin A properties in LAP2α knockout versus wildtype cells, we performed biochemical extraction in buffers containing detergent and 150–500 mM salt. Lamins A and C were significantly less extractable in the absence of LAP2α in both, mouse fibroblasts and HeLa cells (Figure 4C and Figure 4—figure supplement 1A), consistent with a shift of nucleoplasmic lamins into more stable, possibly higher assembly structures or with stronger interactions of lamin complexes with nuclear structures upon loss of LAP2α. Next, we aimed at visualizing potentially higher assembly structures in LAP2α knockout cells by high-resolution microscopy. While microscopy of untreated cells did not reveal clear differences in the appearance of nucleoplasmic lamin structures (Figure 4D, upper panel), extraction of cells in detergent and salt-containing buffers prior to fixation revealed significantly more and larger, extraction-resistant lamin A structures in the nuclear interior in LAP2α knockout versus wildtype mEos3.2-lamin A/C cells (Figure 4D and quantification in Figure 4E). Strikingly, while in wildtype cells these structures appeared as few punctae, LAP2α knockout cells displayed significantly more stable structures of an extended, possibly filamentous appearance (Figure 4D and E).

Altogether, loss of LAP2α does not lead to a reduction of the nucleoplasmic pool of lamin A/C, it rather affects its mobility, interaction and/or assembly state.

LAP2α binds to intranuclear lamin A/C and inhibits its assembly

In order to better understand how LAP2α might influence the mobility and/or assembly state of nucleoplasmic lamin A/C, we performed in vitro interaction analyses of these proteins. Bacterially expressed, purified recombinant lamin A and LAP2α in urea-containing buffer, were dialyzed alone or together into lamin assembly buffer containing 300 mM NaCl, a salt concentration where lamin A does not form higher assembly structures (Foeger et al., 2006), and analyzed the protein samples by sucrose gradient centrifugation. (Figure 5A). While both proteins alone sedimented mostly to fractions 2 and 3 of the sucrose gradient, their sedimentation pattern shifted toward lower fractions in the protein mix, consistent with the formation of larger hetero-complexes (Figure 5A). As a control, a LAP2α mutant lacking the C-terminus that mediates interaction with A-type lamins (Dechat et al., 2000) did not alter the sedimentation pattern of lamin A (Figure 5B). Next, the proteins were dialyzed into buffer with a lower salt concentration (100 mM NaCl) allowing formation of higher assembly lamin structures (Foeger et al., 2006). High-molecular-mass lamin A assemblies were separated from those of lower oligomeric states by centrifugation and analyzed by gel electrophoresis (Figure 5C). Strikingly, the presence of LAP2α reduced the amount of pelleted high-molecular-mass lamin A assemblies by half, indicating that LAP2α impairs lamin A assembly into larger structures (Figure 5C). Thus, binding of LAP2α to lamin A may directly impair the formation of higher order lamin assemblies.

LAP2α binds to lamin A/C and inhibits their assembly without altering lamin phosphorylation.

(A, B). Purified recombinant lamin A and LAP2α full length (f.l.) (A) or lamin A-binding mutant LAP2α1-414 (B) were dialyzed either alone or together into assembly buffer with 300 mM NaCl. Samples were separated on a 10% to 30% sucrose gradient, followed by collection of fractions and quantification of protein bands. Exemplary Coomassie stained gels of fractions ,supernatant’ (SN) to six are shown on the left. The calculated protein amount per fraction (% of total protein) was plotted as curve chart on the right. (C) Lamin A and LAP2α were dialyzed either alone or together into assembly buffer as in (D), but with 100 mM NaCl, enabling formation of higher assembly lamin structures. After centrifugation, total (T), supernatant (SN) and pellet (P) fractions were analyzed on a Coomassie gel, protein bands were quantified and normalized to total protein levels. Graph displays lamin A levels in the pellet fraction of mixed samples (lamin A + LAP2α) as average fold difference ± S.E.M over samples with lamin A alone, n = 5 (single data points are displayed). (D) Lamin A/C was immunoprecipitated from mEos3.2-lamin A/C WT cells and LAP2α and lamin A/C knockout controls using the indicated antibodies recognizing different regions of lamin A/C. Immunoprecipitates were analyzed by western blotting using the indicated antibodies (anti lamin A/C 3A6, anti pS22 lamin A/C, anti LAP2α 1H11). (E) mEos3.2-Lmna WT and LAP2α KO sg1 cells were co-cultured and processed for immunofluorescence microscopy using antibodies specific to lamin A/C phosphorylated at serine 22 (LA/C pS22), antibody 1H11 against LAP2α, and DAPI to visualize DNA. Bar: 10 μm. (F) Isogenic LAP2α KO or WT clones were processed for western blotting using antibodies against lamin A/C phosphorylated at specific residues as indicated or a pan-lamin A/C antibody (E1). Western blot signals for phosphorylated lamins were quantified, normalized to total lamins A and C and expressed as fold difference to the WT samples (graphs on the right). Graphs display average fold difference ± S.E.M. nWT = 3, nKO = 3 (single data points are displayed), p values (paired student’s t-test on log transformed values) are: lamin A pS392 p = 0.602, lamin C pS392 p = 0.493, lamin A pS22 p = 0.841, lamin C pS22 p = 0.826 (non-significant).

To confirm that LAP2α and lamin A/C interact in vivo in mEos3.2-lamin A/C cell lines, we performed co-immunoprecipitation using different lamin A/C antibodies. In accordance with previous data (Dechat et al., 2000; Gesson et al., 2016), antibodies to the lamin A/C N-terminus co-precipitated LAP2α from cell extracts (Figure 5D, antibodies N18 and E1), whereas an antibody to the lamin A/C C-terminus failed to co-precipitate LAP2α (Figure 5D, antibody 3A6), likely because binding of LAP2α to the lamin A/C C-terminus masks the epitope recognized by this antibody (Gesson et a., 2016). Thus, LAP2α interacts with lamin A/C in vivo and may thereby reduce the formation of higher order lamin assemblies in the nuclear interior.

Phosphorylation is another process potentially regulating lamin assembly and mobility, as particularly lamin phosphorylation at serines at position 22 and 392 was described to regulate lamina disassembly during mitosis, as well as lamin A/C localization, mobility and solubility in interphase cells (Heald and McKeon, 1990; Kochin et al., 2014). Thus, we wanted to test whether LAP2α may affect lamin A/C assembly indirectly, by regulating its phosphorylation status. Antibodies specifically recognizing lamin A/C phosphorylated at S22 revealed a similar punctate nucleoplasmic staining with complete absence of peripheral lamina staining in both LAP2α wildtype and knockout cells (Figure 5E and Figure 4—figure supplement 1C; Kochin et al., 2014) and total levels of pS22 lamin A/C and pS392 lamin A/C were unchanged in knockout versus wildtype cells (Figure 5E and F, Figure 4—figure supplement 1B and C). In addition, the relative level of detergent/salt-extractable pS22 lamins A and C were unchanged in LAP2α knockout versus wildtype cells, in stark contrast to the significantly reduced levels of extractable total lamin A/C (Figure 4C). Thus, LAP2α does neither affect the levels nor properties of phosphorylated lamins A and C. However, LAP2α can bind to pS22 lamin A/C as shown by coprecipitation from cell lysates (Figure 5D, antibody pS22). We concluded that LAP2α binds both un(der)phosphorylated and pS22 lamin A/C, but affects preferentially the assembly state of un(der)phosphorylated lamin A/C, keeping it in a mobile, lower assembly state in a phosphorylation-independent manner.

Reduced lamin A/C mobility in the absence of LAP2α decreases chromatin diffusion

To address the relevance of mobile intranuclear lamin A structures for nuclear functions and the consequences upon its loss in LAP2α-deficient cells, we tested chromatin mobility, since intranuclear lamins and LAP2α were found to bind to chromatin in the nuclear interior (Bronshtein et al., 2015; Gesson et al., 2016). Chromatin shows anomalous sub-diffusion, which – unlike free (normal) diffusion – is a motion that is slowed by constraints, such as temporal binding to nucleoplasmic lamins (Bronshtein et al., 2015). Depletion of lamin A/C was shown to increase chromatin motion and to change the type of diffusion of telomeres, centromeres and genomic loci in the nuclear interior toward normal unrestricted diffusion (Bronshtein et al., 2015). To test how depletion of LAP2α affects chromatin motion, we expressed fluorescently labeled TRF1 (to detect telomeres) in mEos3.2-Lmna wildtype and LAP2α knockout cells and analyzed telomere trajectories for 20.5 min to determine the diffusion volume of telomeres (Bronshtein et al., 2015; Vivante et al., 2019). Strikingly, telomere motion was significantly reduced in the absence of LAP2α when compared to wildtype cells (Figure 6A, left panel, see also MSD plot in Figure 6B), and this effect was predominantly dependent on the presence of lamin A/C, since additional depletion of lamin A/C in LAP2α knockout cells (Lmna/Lap2α double knockout, see Figure 6—figure supplement 1) increased the telomere motion volume to levels similar to those observed in Lmna single knockout fibroblasts (Figure 6A , right panel). Thus, the absence of LAP2α and the resulting changes in nucleoplasmic lamin mobility toward a more stable form have direct functional consequences on chromatin diffusion in the nuclear interior, significantly reducing telomere motion.

Figure 6 with 1 supplement see all
Lower lamin A/C mobility in the absence of LAP2α leads to a reduction in telomere movement.

mEos3.2-Lmna WT or mEos3.2-Lmna fibroblasts lacking LAP2α (Lap2α KO sg1), lamin A/C (Lmna KO, see Figure 6—figure supplement 1), or both (Lmna/Lap2α DKO) were transiently transfected with a plasmid expressing DsRed-TRF1 to fluorescently label telomeres. The volume of telomere motion was then calculated based on its trajectory. (A) Box plots compare telomere motion between WT and Lap2α KO sg1 (left; ***p<0.0001 using Mann-Whitney U test, effect size d = 0.62) or between all four genotypes (right; Kruskal-Wallis test p<0.0001; post tests for WT ctrl versus each genotype are all highly significant with ***p<0.0001). The median is depicted within the first and third quartiles; whiskers: minimal and maximal datapoint excluding outliers (see also Figure 6—source data 1). (B) Mean square displacement (MSD) curves for telomere diffusion in mEos3.2-Lmna WT and LAP2α KO sg1 cells. The shaded area shows the distribution of 70% of the population. nWT = 471, nLap2αKO=562, nLmnaKO = 925, nDKO = 1298.

Figure 6—source data 1

Volume of telomere movement in WT, LAP2α KO, Lmna KO and DKO cells.

https://cdn.elifesciences.org/articles/63476/elife-63476-fig6-data1-v2.xlsx

Interestingly, chromatin movement was slightly, but significantly increased in Lmna/Lap2α double knockout cells when compared to Lmna single knockout fibroblasts (Figure 6A, right panel, Kruskal Wallis post test p<0.0001, effect size d = 0.3). This suggests that LAP2α may have additional lamin A/C-independent functions in the regulation of chromatin movement, which is in accordance with the broad, genome-wide interaction of LAP2α with euchromatin (Gesson et al., 2016). Thus, LAP2α may affect chromatin movement in the nuclear interior by at least two different means: (1) by regulating lamin A/C assembly and/or chromatin interaction, thereby affecting lamin-A-mediated chromatin regulation and (2) by binding to chromatin and causing a slight, but significant reduction in chromatin motion in the nuclear interior.

In summary, our data provide novel insight into the formation and regulation of nucleoplasmic lamin A/C by LAP2α (Figure 7). While the nucleoplasmic pool of lamin A/C is formed independently of LAP2α, the properties of lamin A/C in the nuclear interior are significantly affected by LAP2α in a lamin serine 22 phosphorylation-independent manner. Binding of LAP2α impairs formation of higher order structures of lamin A/C and/or their interaction with chromatin, thereby keeping them in a mobile and low assembly state, which in turn affects chromatin mobility.

Model of LAP2α-dependent regulation of A-type lamins in the nucleoplasm.

LAP2α dynamically interacts with lamins A and C in the nucleoplasm, independent of their phosphorylation status. While lamins at the periphery interact mainly with heterochromatic lamina-associated domains (LADs), nucleoplasmic A-type lamins and LAP2α bind to euchromatic genome regions regulating chromatin mobility. When hypophosphorylated lamins are not bound to LAP2α, they form larger, stable assemblies in the nucleoplasm. In the absence of LAP2α, these lamin A/C structures become more frequent, leading to altered, putatively less dynamic lamin A/C chromatin interaction and slower chromatin movement.

Discussion

Here, we show that LAP2α, a known binding partner of nucleoplasmic lamin A/C (Dechat et al., 2000; Naetar and Foisner, 2009) is essential to maintain lamins A and C in the nuclear interior in a mobile and low assembly state. This occurs in a lamin A/C S22 phosphorylation-independent manner, likely by inhibiting lamin assembly through direct interaction of LAP2α with lamin A/C and/or by changing interactions of nucleoplasmic lamin complexes with chromatin.

Our study confirms and extends the finding that lamins A and C in the nuclear interior have fundamentally different properties compared to their counterparts at the nuclear periphery. While lamins at the nuclear periphery form stable 3.5-nm-wide filaments (de Leeuw et al., 2018; Moir et al., 2000; Shimi et al., 2008; Turgay et al., 2017), lamins in the nuclear interior are highly mobile (Broers et al., 1999; Bronshtein et al., 2015; Shimi et al., 2008) and easily extractable in salt and detergent-containing buffers (Kolb et al., 2011; Naetar et al., 2008). However, very little is known about the mechanisms securing these unique properties of lamins in the nuclear interior, given that in solution they have a strong drive to assemble and tend to form higher order structures (Aebi et al., 1986; Moir et al., 1991). Immunofluorescence microscopy has previously shown that intranuclear lamin A/C staining is significantly reduced in cells and tissues from LAP2α knockout mice (Naetar et al., 2008), leading to a model, where LAP2α is required for the formation and/or maintenance of the nucleoplasmic lamin pool (Naetar and Foisner, 2009). Based on data shown in this study, we propose a modified and updated version of the model suggesting that LAP2α is essential to maintain the highly mobile and soluble state of lamins in the nuclear interior. In the absence of LAP2α, lamins in the nucleoplasm are not lost but they are transformed into more stable, immobile structures (Figure 7) that can no longer be recognized by lamin A/C antibodies directed to an N-terminal epitope, which – in wildtype cells – favors nucleoplasmic over peripheral lamin staining (Gesson et al., 2016). These findings can be explained by the formation of higher order lamin complexes in the nuclear interior, whose properties may resemble more those of lamins at the nuclear periphery. This is supported by the increased resistance of nucleoplasmic lamins to extraction with detergent and salt-containing buffers and their decreased mobility in the absence of LAP2α. Moreover, high-resolution microscopy revealed significantly more and, importantly, larger extraction-resistant structures in LAP2α knockout versus wildtype cells. As for potential mechanisms describing how LAP2α can maintain lamins in a soluble and dynamic state, we can envisage at least two different not mutually exclusive mechanisms. Our in vitro data using purified recombinant LAP2α and lamin A protein suggest that direct binding of LAP2α to lamins may prevent them from assembling into higher order structures. However, one has to keep in mind that the in vitro assembly of lamins may not fully represent lamin assembly pathways within a cell, as previously indicated in a study using lamin-binding designed ankyrin repeat proteins (DARPins), where DARPins inhibiting in vitro lamin A assembly had no impact on incorporation of lamin A into the lamina in vivo and, vice versa, DARPins affecting the incorporation of lamin A into the nuclear lamina in vivo had no effect on lamin assembly in vitro. Nevertheless, we showed in a previous study that, unlike wildtype lamin A/C staining (Naetar et al., 2008 and Figure 3), antibody-staining of the assembly-deficient lamin AΔK32 mutant in the nuclear interior is not affected in LAP2α knockout cells (Pilat et al., 2013). This suggests that nucleoplasmic lamin A assembly in the absence of LAP2α leads to the observed antibody epitope masking.

Besides affecting lamin assembly, LAP2α may also regulate the interaction of A-type lamin complexes with chromatin. In this model, the absence of LAP2α would lead to a tighter and more stable association of lamins with chromatin, which would in turn also lead to an increased extraction resistance and possibly decreased mobility. In support of this model, the post-fixation digestion of formaldehyde-fixed cells with DNase/RNase fully recovered the intranuclear lamin A/C staining using antibodies to the lamin A/C N-terminus, and chromatin motion was slowed down upon LAP2α knockout in a lamin A/C-dependent manner. However, the finding that the intranuclear lamin structures are resistant to 500 mM high salt, which is expected to extract chromatin-associated proteins (Herrmann et al., 2017), supports the notion that larger lamin A structures can be formed independently of chromatin. As these different scenarios are not mutually exclusive, we favor a model in which the absence of LAP2α induces the formation of higher order lamin A assemblies, which in turn may bind more tightly and stably to chromatin (Figure 7).

As phosphorylation of lamins provides another means to regulate lamin filament (dis)assembly in mitosis (Heald and McKeon, 1990; Ward and Kirschner, 1990) and to influence their assembly state, solubility and mobility in interphase (Buxboim et al., 2014; Kochin et al., 2014; Moiseeva et al., 2016), we tested the relationship between lamin phosphorylation and interaction with LAP2α. We found that LAP2α loss does neither affect total levels of lamins phosphorylated at serines 22 and 392 nor does it change the localization and solubility of phosphorylated lamins. Vice versa, LAP2α can interact with both phosphorylated and un(der)phosphorylated lamins, but it affects only the solubility and assembly state of un(der)phosphorylated lamins. Thus, interaction with LAP2α and lamin phosphorylation seem to be two independent, non-mutually exclusive mechanisms for keeping lamins in a low assembly state.

Surprisingly, the initial formation of the nucleoplasmic lamin A/C pool does not require LAP2α, as it can form normally from newly synthesized pre-lamin A and from soluble mitotic lamins in LAP2α knockout cells. Thus, we propose that these processes may primarily depend on other mechanisms, such as lamin A/C phosphorylation. At the onset of mitosis, lamins are targeted by cyclin-dependent kinase 1 (Cdk1) particularly at serines 22 and 392 in the N-terminal head and proximal to the C-terminal end of the rod, respectively (Ward and Kirschner, 1990), leading to a complete disassembly of the lamina (Dechat et al., 2004; Heald and McKeon, 1990; Moir et al., 2000). During post-mitotic nuclear envelope reassembly, lamins A and C are imported into the nucleoplasm of newly formed nuclei, followed by gradual dephosphorylation and re-assembly at the nuclear periphery (Moir et al., 2000; Steen et al., 2003; Steen and Collas, 2001). It is tempting to speculate that the majority of lamins A and C is still phosphorylated at S22 and possibly other residues when they are re-imported into the nucleoplasm after post-mitotic nuclear membrane reassembly, preventing their incorporation into the peripheral lamina. In the nucleoplasm, a fraction of the phosphorylated lamins may then bind to LAP2α. When lamins are gradually dephosphorylated during G1 (Steen et al., 2003; Steen and Collas, 2001), LAP2α-bound lamin A/C may be maintained in a low assembly state, while free lamin A/C can assemble into the lamina at the periphery. Thus, in the absence of LAP2α, the nucleoplasmic lamin A/C pool may still form, but, whereas phosphorylated lamins stay soluble independently of LAP2α, hypophosphorylated lamins tend to form higher-order structures at the ‘wrong’ position, that is within the nucleoplasm as opposed to the nuclear periphery in LAP2α knockout cells. This model suggests that in wild-type cells, there are three pools of intranuclear A-type lamins (Figure 7): (1) lamins phosphorylated at S22 (and possibly other residues) that are soluble independent of LAP2α; (2) hypophosphorylated lamins bound to LAP2α that are soluble independent of their phosphorylation status; and (3) hypophosphorylated lamins not bound to LAP2α that tend to form immobile and extraction-resistant structures in the nuclear interior. Previous reports and our own data showing that a small fraction of nucleoplasmic A-type lamins displayed low mobility (Bronshtein et al., 2015; Shimi et al., 2008) in wildtype cells supports this model. In the absence of LAP2α, the balance between these different intranuclear lamin structures is disturbed toward increased levels of immobile, higher order lamin A/C structures.

What is the physiological relevance of a dynamic lamin A/C pool in the nucleoplasm and how is the nucleoplasmic lamin A/C pool regulated during various cellular processes? We propose that the unique dynamic properties allow intranuclear A-type lamins to fulfill a set of functions that is different from those covered by the peripheral lamina. One of the most prominent examples for different functions of the peripheral versus intranuclear lamins is their different role in chromatin organization. While the peripheral lamina is well known to mediate stable interaction with heterochromatic genomic regions, termed lamina-associated domains (LADs) (Kind et al., 2013; van Steensel and Belmont, 2017), thereby contributing to gene silencing in general (Leemans et al., 2019) or during differentiation (Robson et al., 2016), lamins in the nuclear interior have a much more complex role in regulating open chromatin and gene expression. Lamin A/C in the nuclear interior interact primarily with open euchromatic regions of the genome (Gesson et al., 2016; Lund et al., 2015) and affect their epigenetic landscape (Bianchi et al., 2020; Briand et al., 2018; Cesarini et al., 2015; Oldenburg et al., 2017). Furthermore, lamin A/C in the nucleoplasm has also been shown to directly interact with promoters and enhancers and seems to be involved in both positive and negative gene regulations (Ikegami et al., 2020; Lund and Collas, 2013; Lund et al., 2013). It is conceivable that gene regulatory pathways have to be highly flexible and dynamic to respond efficiently to internal and external cues and thus require a dynamic lamin complex that can dynamically associate with and regulate chromatin. Indeed, loss of LAP2α (Gesson et al., 2016) or disease-linked mutations in lamins (Bianchi et al., 2020; Briand et al., 2018; Ikegami et al., 2020; Oldenburg et al., 2017) affect epigenetic regulation and/or gene expression.

Another function of nucleoplasmic lamins is to regulate the mobility of chromatin in the three-dimensional nuclear space. Dynamic interactions of lamin complexes with telomeres and other genomic loci has been shown to limit free diffusion of chromatin inside the nucleus (Bronshtein et al., 2015). We demonstrate here that knockout of LAP2α reduces telomere motion mostly in a lamin A/C-dependent manner, most likely through the formation of more stable lamin A/C structures that bind chromatin more tightly and stably. It is reasonable to assume that a tightly regulated movement of genes inside the nucleus is important for controlling coordinated regulation of gene expression during various cellular processes. Notably, LAP2α may have another lamin A-independent role in the regulation of chromatin diffusion. By direct binding to chromatin, LAP2α may additionally slow down chromatin diffusion. However, the major contribution to restricted, anomalous chromatin diffusion in the nuclear interior seems to stem from nucleoplasmic lamin A/C, as its depletion has the highest impact on chromatin movement.

Overall, we speculate that a dynamic nucleoplasmic lamin A/C complex may be required to maintain genomic plasticity, allowing the cell to efficiently respond to external and internal cues, such as during cell differentiation. In line with this hypothesis, we found nucleoplasmic lamins and LAP2α in proliferating tissue progenitor cells in the epidermis, colon, skeletal muscle and in preadipocytes, while upon their differentiation (Dorner et al., 2006; Gotic et al., 2010; Markiewicz et al., 2005; Naetar et al., 2008) or cell cycle exit into a quiescence or senescence state (Naetar et al., 2007) LAP2α was downregulated at transcriptional and protein level leading to loss of the dynamic nucleoplasmic lamin A/C pool. Thus, when genome plasticity is no longer required, such es in terminally differentiated cells, dynamic lamin A/C levels may be reduced through downregulation of LAP2α. Accordingly, the absence of the dynamic nucleoplasmic lamin A/C pool in the LAP2α knockout progenitor cells leads to impaired or delayed differentiation (Gotic et al., 2010; Naetar et al., 2008). This regulatory pathway may also be impaired in lamin-linked diseases, such as the premature aging disease Hutchinson Gilford Progeria Syndrome (HGPS), where LAP2α expression is lost, leading to premature senescence, while re-expression of LAP2α in HGPS cells rescues cell proliferation (Vidak et al., 2015).

In summary, we propose that lamin A/ C complexes in the nuclear interior are in a dynamic state, established by LAP2α, which is a prerequisite for their multiple roles in chromatin regulation both on a genome-wide scale and in a gene-specific manner during cell differentiation.

Materials and methods

Key resources table
Reagent type (species)
or resource
DesignationSource or referenceIdentifiersAdditional information
Gene (Mus musculus)LmnaGenBankGene:ID_16905
Gene (M. musculus)Tmpo; Lap2GenBankGene:ID_21917
Gene (Homo sapiens)TMPO; LAP2GenBankGene:ID_7112
Strain, strain
background (E.c oli)
NEB5-alpha Competent
E. coli (high efficiency)
New England BiolabsCat# C2987Chemically competent
Strain, strain
background (E. coli)
BL21-CodonPlus (DE3)PMID:10562549
Cell line (M. musculus)mEos3.2-Lmna wildtype
dermal fibroblasts
This paperSee Materials and methods, section ‘Generation and cultivation of cell lines’
Cell line (M. musculus)mEos3.2- Lmna LAP2α
knockout fibroblasts
This paperSee Materials and methods, section ‘Generation and cultivation of cell lines’
Cell line (M. musculus)mEos3.2- Lmna lamin A/C
knockout fibroblasts
This paperSee Materials and methods, section ‘Generation and cultivation of cell lines’
Cell line (M. musculus)mEos3.2- Lmna LAP2α-lamin
A/C double knockout fibroblasts
This paperSee Materials and methods, section ‘Generation and cultivation of cell lines’
Cell line (H. sapiens)LAP2α knockout HeLa cellsThis paperSee Materials and methods, section ‘Generation and cultivation of cell lines’
Transfected construct (M. musculus)Lmna-targeting constructThis paperSee Materials and methods, section ‘Generation of Lmna-specific targeting construct’
Transfected construct
(M. musculus and H. sapiens)
sgRNAs targeting Lmna
or Lap2 in pX458
This paperSee Materials and methods, section ‘Generation and cultivation of cell lines’
AntibodyAnti-LAP2α
(rabbit, polyclonal)
PMID:11864981WB (1:5000)
IF (1:800)
AntibodyAnti-LAP2α 1H11
(mouse, monoclonal)
Max Perutz Labs Monoclonal Antibody facility, see also PMID:26798136WB (1:100)
IF (1:30)
AntibodyAnti-LAP2α Ab15-2
(mouse, monoclonal)
PMID:9707448WB (undiluted hybridoma supernatant)
AntibodyAnti-LAP2 Ab12
(mouse, monoclonal)
PMID:9707448WB (1:4)
AntibodyAnti-lamin A/C 3A6
(mouse, monoclonal)
Max Perutz Labs Monoclonal Antibody facility, see also PMID:26798136WB (1:50)
IF (1:50)
AntibodyAnti-lamin A/C E1
(mouse, monoclonal)
Santa Cruz BiotechnologyCat# sc-376248,
RRID: AB_10991536
WB (1:1000)
IF (1:100)
AntibodyAnti-lamin A/C
N18 (goat, polyclonal)
Santa Cruz BiotechnologyCat# sc-6215,
RRID:AB_648152
WB (1:1000)
IF (1:100)
AntibodyAnti-lamin A/C pS22
(rabbit polyclonal)
Invitrogen/ Thermo Fisher ScientificCat# PA5-17113,
RRID: AB_10989809
WB (1:200)
IF (1:50)
AntibodyAnti-lamin A/C pS22 D2B2E
(rabbit, monoclonal)
Cell SignalingCat# 13448,
RRID:AB_2798221
IP (5 μl/IP)
AntibodyAnti-lamin A/C pS392 (rabbit, polyclonal)AbcamCat# ab58528,
RRID:AB_883054
WB (1:500)
AntibodyAnti-lamin B1 (rabbit, polyclonal)ProteintechCat# 12987–1-AP,
RRID: AB_2136290
WB (1:500)
IF (1:100)
AntibodyAnti-mEos2 (rabbit polyclonal)BadrillaCat# A010-mEOS2,
RRID: AB_2773027
IF (1:200)
AntibodyAnti-γ-tubulin GTU88
(mouse, monoclonal)
Sigma AldrichCat# T6557,
RRID:AB_477584
WB (1:5000)
AntibodyAnti-actin I-19 (goat, polyclonal)Santa Cruz BiotechnologyCat# sc-1616,
RRID:AB_630836
WB (1:500)
Recombinant DNA reagentpSpCas9(BB)−2A-GFP;
pX458 (plasmid)
AddgeneRRID:Addgene_48138sgRNAs targeting Lmna or Lap2 were inserted into this vector
Recombinant DNA reagentpSpCas9(BB)−2A-mCherry (plasmid)This paperSee Materials and methods, section ‘Vectors’
Recombinant DNA reagentpEGFP-myc-Lamin A (plasmid)This paperSee Materials and methods, section 'Vectors'
Recombinant DNA reagentdsRed-TRF1 (plasmid)PMID:26299252
Sequence-based reagenthLAP2α sgRNA1This papersgRNAAGTTCGGTACTGCCCAAAGG
Sequence-based reagenthLAP2α sgRNA2This papersgRNAGGAACTACTCCCTCTGGTGG
Sequence-based reagentmLmna sgRNA1This papersgRNACACTCGGATCACCCGGCTGC
Sequence-based reagentmLmna sgRNA2This papersgRNAACGCACGCGATCGATGTACA
Sequence-based reagentmLap2α sgRNA1This papersgRNAAGTTCAGTCCTGCCCAAAGG
Sequence-based reagentmLap2α sgRNA3This papersgRNACAAGAAAGTGAAGTCCGCTA
Peptide, recombinant proteinQ5 High Fidelity DNA polymeraseNew England BiolabsCat# M0491
Commercial assay or kitRevertAid First Strand cDNA synthesis kitThermo Fisher ScientificCat# K1622
Commercial assay or kitKAPA SYBR Green 2x PCR master mixKapa BiosystemsCat# KK4602
Commercial assay or kitGibson assembly master mixNew England BiolabsCat# E2611
Commercial assay or kitGoTaq green master mixPromegaCat# M7122
Software, algorithmTIDEPMID:25300484https://tide.nki.nl/
Software, algorithmVisiview 4.4 softwareVisitron systems
Software, algorithmFIJIOpen sourceRRID:SCR_002285https://imagej.net/Fiji
Software, algorithmZeiss ZEN 2.1 softwareZeiss
Software, algorithmMatlabMathWorksRRID:SCR_001622
Software, algorithmSymPho Time softwarePicoQuant

Generation and cultivation of cell lines

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To generate mEos3.2-Lmna mouse dermal fibroblast cell lines, immortalized fibroblasts derived from the back skin of wild-type and LAP2α knockout littermates (Naetar et al., 2008) were transfected with the vector pSpCas9(BB)−2A-GFP (pX458, plasmid #48138 from Addgene, Watertown, MA) (Ran et al., 2013) carrying mLmna-specific sgRNA1 (see Key Resource Table) and Cas9 from S. pyogenes with 2A-EGFP, and a Lmna-specific targeting construct, where the fluorophore mEos3.2 (Zhang et al., 2012) was inserted in frame into exon 1 of the Lmna gene in front of the first codon (Figure 2A, see also ‘Generation of Lmna-specific targeting construct’). Cells were sorted for EGFP expression using a FACS Aria Illu (Becton Dickinson, Franklin Lakes, NJ), cultivated for 14 days, followed by a single cell sort for mEos3.2-expressing cells. Single cell clones were outgrown and further characterized by genotyping PCR, long-range PCR, sequencing of the targeted locus and Western blotting.

The wildtype mEos3.2-Lmna clone #21 was selected to create isogenic Lap2α knockout clones, as well as Lmna knockout and Lmna/Lap2α double knockout clones. Cells were transfected with the vector pSpCas9(BB)−2A-mCherry (modified pSpCas9(BB)−2A-GFP, see also ‘Vectors’) carrying mLap2α-specific sgRNA1 or 3, or mLmna-specific sgRNA2 (see Key Resource Table) and Cas 9 from S. pyogenes with 2A-mCherry. mCherry-positive single cells were FACS-sorted and knockout clones were identified by western blot. Knockout clones and wildtype control clones were further characterized by sequencing of a PCR product derived from isolated genomic DNA spanning the expected Cas9 cut site (primers mLap2α-f and -r, or mLmna-f and -r: see ‘Primers’). Sequences were analyzed using the TIDE software available online (https://tide.nki.nl/) (Brinkman et al., 2014). Absence of continuous Cas9 expression was verified by mCherry FACS analysis using an LSR Fortessa (Becton Dickinson).

To generate LAP2α knockout HeLa cell clones, HeLa cells were transfected with pSpCas9(BB)−2A-GFP carrying hLAP2α-specific sgRNA1 or 2 (see Key Resource Table). GFP-positive cells were sorted and plated for generation of single cell clones. LAP2α knockout clones were identified by western blotting. Knockout clones and wildtype control clones were further characterized by sequencing of a PCR product spanning the expected Cas9 cut site (primers hLAP2α-f and -r: see ‘Primers’). Sequences were analyzed using TIDE. Absence of continuous Cas9 expression was verified by GFP FACS analysis.

All cells were maintained at 37°C and 5% CO2 in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal calf serum (FCS), 2 mM glutamine, 50 U/ml penicillin and 50 μg/ml streptomycin (P/S) (all from Sigma-Aldrich, St. Louis, MO). Non-essential amino acids (PAN-Biotech, Aidenbach, Germany) were routinely added for mouse dermal fibroblasts. After sorting, cells were kept in medium additionally containing Antimycotic-Antibiotic (Gibco/Thermo Fisher Scientific, Waltham, MA) and Plasmocin (25 μg/ml, Invivogen, San Diego, CA) to prevent sorter-induced contamination. Cells were routinely inspected for mycoplasma contamination after staining nuclei with DAPI or Hoechst dye (Thermo Fisher Scientific). Additionally, especially if mycoplasma contamination was suspected based on DAPI staining, cells were tested for mycoplasma contamination using the MycoAlert Mycoplasma detection kit (Lonza, Basel, Switzerland). To determine cell viability, cells were lysed in CASYblue (OLS OMNI Life Science, Bremen, Germany), containing MeOH. The lysed cell sample was measured using a Casy Counter (OLS OMNI Life Science) and the cell volume profile was used to define the volume cut-off for dead cells. Living cell samples were then measured according to manufacturer’s instructions using these settings to determine the percentage of viable cells within the sample.

Live cell imaging

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Cells were plated on 35 mm glass-bottom μ-dishes (Ibidi, Gräfelfing, Germany) in high-glucose phenol-red free DMEM (Gibco Fluorobrite) supplemented with 10% FCS, L-glutamine, and P/S. For mEos3.2 cells, dishes were pre-coated with Cell-tak (Corning, Corning, NY) according to manufacturer’s instructions. HeLa cells were transfected with plasmids expressing EGFP-tagged pre-lamin A, mature lamin A, pre-lamin A delK32 or mature lamin A delK32 (see’Vectors’) using nanofectin (PAA Laboratories, Toronto, Canada) according to manufacturer’s instructions. To avoid substantial overexpression, a maximum of 1 μg plasmid DNA per 35 mm dish was used. Cells were imaged 5 hr or 24 hr post transfection.

Imaging was performed using a Visitron Spinning disc microscope (Zeiss, Oberkochen, Germany) under controlled environmental conditions at 37°C and 5% CO2, using a Plan-Apochromat 63x/1.4 Oil DIC III objective and an EM-CCD camera. Images (Z stacks) were obtained by automated multipositioning (40 positions per 35 mm dish) every 20 min with the same excitation strength and exposure time using Visiview 4.4 software (Visitron Systems, Puchheim, Germany). Images were processed in FIJI. For experiments with HeLa cells, all time-lapse images were smoothed by 1-pixel radius Gaussian Blur filter, the frame was cropped to fit the cells of interest and substacks were created from Z stacks to include the focal plane of appropriate time points starting with the beginning of mitosis until late G1. These substacks were then used to create short movies and the images presented in the results section. The data were quantified manually by drawing a three pixel wide bar across the nuclei of selected cells and creating plot profiles for each cell. The peripheral (P) lamin signal was calculated by averaging two peak measurements, one from each end of the drawn line, and the nucleoplasmic (N) intensity as the average of 30 measurements spanning the center of the drawn line, followed by calculation of the N/P ratio.

Automated quantification of the N/P ratio for mEos3.2-lamin A/C cells was done using custom-made FIJI plugins, where mitotic cells are manually extracted from the images, followed by automatic analysis of time series stacks and data visualization of nucleoplasmic to peripheral signal intensity ratio over time. Specifically, frames, slices and fields of view with relevant cells were manually selected and extracted for automated analysis. Nuclei were segmented by thresholding, marking regions of interest. These regions were further reduced and subtracted to define nucleoplasm and nuclear periphery. Average intensity values of mEos3.2 were extracted from these regions and the signal ratio (nucleoplasm/periphery) was calculated.

Immunofluorescence and cell cycle staging using DAPI

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For standard immunofluorescence (IF), cells were seeded on uncoated glass coverslips (1.5H, Marienfeld-Superior, Lauda-Königshofen, Germany) and fixed with 4% paraformaldehyde for 10 min at room temperature. To stop fixation and permeabilize the cells, coverslips were incubated in PBS with 0.1% Triton X-100 and 50 mM NH4Cl, followed by incubation in primary and secondary antibody (for a list of all antibodies see Key Resource Table). To reverse chromatin-dependent epitope masking of nucleoplasmic lamins, cells were treated with 100 μg/ml DNase I and 100 μg/ml RNase A for 30 min at room temperature. DNA was stained with DAPI (0.5 μg/ml) and cells were mounted using Vectashield (Vector Laboratories, Burlingame, CA). Immunofluorescence slides were routinely imaged using an LSM710 confocal microscope (Zeiss) equipped with a Plan-Apochromat 63x/1.4 Oil DIC M27 objective and standard photomultiplier tubes (PMTs) for sequential detection, as well as an Airyscan detector for high-resolution imaging. Image acquisition was done using Zeiss ZEN 2.1 software, followed by image processing using FIJI, including adjustment of the digital offset for high resolution Airy scan images to avoid over- and undersaturation of the fluorescent signal. IF images with standard resolution were smoothened by 1-pixel radius Gaussian Blur filter. Extraction-resistant lamin A/C structures were quantified using a custom-made FIJI plugin, where cells were first defined using the FIJI built-in auto-threshold function on average Z projections of mEos3.2-lamin A/C, followed by identification and quantification (number and area) of intranuclear structures using the threshold function with the built-in ‘Huang’ algorithm, followed by the particle analyzer function, including particles between 0.0001 and 5 μm, but excluding a rim of 0.8 μm from the nuclear periphery to avoid counting of signals within the nuclear lamina.

For DAPI-based cell cycle staging and combined determination of nucleoplasmic to peripheral ratio of mEos3.2-tagged lamin A/C, cells were imaged using the Visitron Spinning disc microscope (Zeiss) and a custom-made slide scanner, allowing the automated acquisition of multiple image stacks (400 fields of view/sample). Images were analyzed using a custom-made FIJI plugin, where the cell cycle stage of each cell was determined based on the DAPI intensity and a cell cycle profile was created. Initial regions of interest were extracted by segmenting nuclei using thresholding and watersheding. DAPI intensity was measured by summing the integrated density of DAPI signals within regions of interest. Initial regions of interest were further reduced and subtracted to define nucleoplasm and nuclear periphery. Intensity of the nucleoplasm was measured by taking the median value of the minimum projection of the regions of interest. Intensity of the nuclear periphery was measured by taking the upper quartile of the maximum projection of the regions of interest. The nucleoplasmic/peripheral lamin A/C ratio of the cells was calculated and plotted against the cell cycle stage.

Continuous photobleaching and fluorescence correlation spectroscopy

Continuous photobleaching was performed using a confocal microscope (Leica TCS SP8 SMD, Leica Microsystems, Wetzlar, Germany). For illumination, the PicoQuant Picosecond Pulsed Diode Laser Head of 40 MHz 470 nm was used (laser intensity was set to ~1 µW). The light is focused onto a small confocal volume through a 63x water immersion objective lens with NA = 1.2 (Leica HC PL APO 63x/1.20 W CORR). The signal is detected by sensitive detectors (Leica HyD SMD) within the detection window (500–600 nm). A specific point was chosen in the nuclear interior and a ‘point measurement’ was performed for measuring the intensity at a high frequency (~1 KHz) for approximately 60 s per cell. CP data analysis was performed with Matlab and a custom-made algorithm (Bronshtein et al., 2015).

Fluorescence correlation spectroscopy (FCS) curves were extracted from the region where the fluorescence intensity is constant (approximately 30 s after beginning of the measurement) in order to avoid inaccuracy due to bleaching. The FCS analysis was done by SymPho Time software (PicoQuant, Berlin, Germany). The best fit was achieved with the FCS Triplet 3D fitting model:

(1) Gt=1+j=0nTrip-1Tjexp-tτTripj-1i=0nDiff-1 ρi1+tτDiffiai1+tτDiffiaik20.5 +GInf
w0=VEffk13π-0.5
Dk=w024τDiff[k]

Model parameters

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  • Number of triplet (dark) states nTrip=1

  • Number of independently diffusing species nDiff=2

  • Effective excitation volume VEff=0.267[fl]

  • Length to diameter ratio of the focal volume k=7.92

  • Anomaly parameter of the ith diffusing species a1,2=1

Fitting parameters

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  • Contribution of the ith diffusing species - ρi

  • Diffusion time of the ith diffusing species - τDiffi

  • Dark (triplet) fractions of molecules - T

  • Lifetime of the dark (triplet) states - τTrip

  • Correlation offset - GInf

  • Effective lateral focal radius at 1e2 intensity - w0

  • Diffusion constant of the kth diffusing species - Dk

To explain the observed differences in the measured diffusion of free lamin A complexes between wild-type and LAP2α knockout cells, the diffusion was modeled making several basic assumptions on protein shape and structure, based on the Stokes-Einstein relation for normal diffusion:

D = κB T6πηα

where kB is the Boltzmann constant, T is the temperature, η is the nucleoplasm viscosity and α is the protein radius, assuming a spherical shape.

D 1α

and one can assume that the protein volume is proportional to its molecular weight. (Erickson, 2009).

Therefore, DWTDKO=αKOαWT=MKOMWT3 , and the ratio of the FCS diffusion rates DWT/DKO should equal the cube root of the inverted nucleoplasmic lamin complex molecular weight ratio MKO/MWT, allowing to make conclusions on the relative molecular weight of measured complexes in WT and KO cells.

Biochemical extraction experiments

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Cells were washed twice with PBS (Sigma-Aldrich) and directly scraped off the plate in cold extraction buffer on ice (20 mM Tris-HCl pH7.5, 150 mM NaCl, 2 mM EGTA, 2 mM MgCl2, 0.5% NP-40, 1 mM DTT, 1 U/ml Benzonase from Novagen/EMD Millipore, Temecula, CA, 1x Complete Protease Inhibitor Cocktail from Sigma-Aldrich, 1x Phosphatase inhibitor cocktail 2 and 3 from Sigma-Aldrich). For mouse fibroblasts, extra NaCl was added to a final concentration of 500 mM. Extracts were incubated for 10 min on ice and then centrifuged for 10 min at 4000 rpm in a Megafuge 1.0R (Haereus, Hanau, Germany). Pellets were resuspended in equal volumes of extraction buffer and sonicated with a Bandelin Sonopuls HD200 sonicator (Bandelin, Berlin, Germany; settings: MS73/D at 50% intensity) for 3 s to solubilize insoluble material. Total, supernatant and pellet fractions were analyzed by immunoblotting.

For in situ extraction of mEos3.2-lamin A/C mouse fibroblasts, cells were grown on glass coverslips (1.5H, Marienfeld) that were pre-coated with Cell-tak (Corning) according to manufacturer’s instructions to avoid detachment of cells due to extraction. Cells were washed twice with PBS and incubated for 10 min in extraction buffer containing 500 mM NaCl (without benzonase), followed by fixation in 4% paraformaldehyde according to the routine IF protocol (see ‘Immunofluorescence and cell cycle staging using DAPI’). To amplify the mEos3.2 signal, cells were stained with an anti-mEos2 antibody (Badrilla, Leeds, UK) following the standard IF protocol.

Recombinant protein expression and sedimentation assays

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Recombinant full-length human LAP2α and the truncation mutant LAP2α1-414 were expressed in E. coli BL21 (DE3) using the plasmid pET23a(+) (Vlcek et al., 1999), which adds a 6x histidine-tag to the C-terminus of the expressed proteins for further affinity purification. Protein expression was induced in bacterial cultures at an OD600nm of 0.6–0.7 with 0.5 mM Isopropyl-β-D-thiogalactopyranoside (IPTG) for 3 hr. Bacterial pellets were then resuspended in 10 mM Tris-HCl pH 8.0, 100 mM NaCl, and 1 mM DTT, followed by cell lysis in the presence of 1x protease inhibitor cocktail (Sigma Aldrich). 5 μg/ml DNase I and 10 μg/ml RNase A were added and inclusion bodies of the respective recombinant proteins were pelleted by centrifugation. Poly-histidine-tagged recombinant proteins were purified using Ni-NTA-Agarose (Qiagen, Hilden, Germany) beads according to manufacturer’s instructions and were analyzed by SDS-polyacrylamide gel electrophoresis (SDS-PAGE). Pure protein fractions were dialyzed twice against 8 M Urea, 10 mM Tris-HCl pH 8.0, 300 mM NaCl and 1 mM DTT using a cut-off of 12–14 kDa to remove Imidazole and ß-Mercaptoethanol. Purified recombinant human wild-type lamin A was a kind gift of Prof. Harald Herrmann, DKFZ, Heidelberg, Germany.

For sucrose density gradient centrifugation, recombinant proteins were dialyzed against 10 mM Tris-HCl pH 7.5, 1 mM DTT, 300 mM NaCl and 10% sucrose and were layered on top of a sucrose gradient ranging from 10% to 30% atop a 70% sucrose cushion in a centrifuge tube. Samples were centrifuged for 20 hr at 190,000 x g in a SW-40 Rotor (Beckman Optima L-70 or Beckman Optima L-80 XP, Beckman Coulter, Brea, CA) at 4°C. Collection of the sedimented protein layers was done by pipetting off fractions from the top to the bottom of the gradient, starting with the applied dialyzed protein sample (supernatant). Aliquots of all gradient fractions were analyzed on an SDS-polyacrylamide gel stained with Gel CodeTM Blue Safe Protein Stain (Thermo Fisher Scientific) and density measurement of the protein bands was done with ImageJ software.

To induce lamin assembly, recombinant proteins were dialyzed stepwise against 10 mM Tris-HCl pH 7.5, 1 mM DTT containing decreasing salt concentrations, starting at 300 mM NaCl, followed by 200 mM NaCl, and ending at 100 mM NaCl. Each step was done at room temperature for 30 min. Samples were then analyzed by centrifugation at 13,000 rpm for 10 min (Eppendorf table top centrifuge, Eppendorf, Hamburg, Germany) at room temperature. Supernatant and pellet fractions were analyzed by SDS-PAGE as described above for sucrose gradient fractions.

Immunoprecipitation and immunoblotting

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For co-immunoprecipitation (IP), cells were scraped off the plate in IP buffer containing 20 mM Tris-HCl pH7.5, 150 mM NaCl, 2 mM EGTA, 2 mM MgCl2, 0.5% NP-40, 1 mM DTT, 1 U/ml Benzonase (Novagen), 1x Complete Protease Inhibitor cocktail (Sigma-Aldrich) and Phosphatase inhibitor cocktail 2 and 3 (Sigma-Aldrich), and incubated 10 min on ice. The soluble fraction after centrifugation for 10 min at 4000 rpm in a Megafuge 1.0R (Haereus) was used as input for the IP (1 mg/IP). Incubation with antibody (5 μg/IP) was done overnight at 4°C, followed by incubation with BSA-blocked proteinA/G dynabeads (Pierce/Thermo Fisher Scientific) for 4 hr at 4°C. Beads were washed three times in IP buffer without benzonase, followed by elution of complexes from the beads using SDS PAGE sample buffer. Samples were analyzed by SDS PAGE and immunoblotting.

Immunoblots were treated with primary antibodies (see Key Resource Table) overnight and with horse radish peroxidase-coupled secondary antibodies (Jackson ImmunoResearch, Westgrove, PA) for 1 hr at room temperature. Signal detection was done using SuperSignal West Pico plus chemiluminescent substrate (Pierce/Thermo Fisher Scientific) and the ChemiDoc Gel Imaging system (Bio-Rad, Hercules, CA). Image analysis and quantification was done using the Image Lab software (Bio-Rad).

Telomere tracking

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Cells were transfected with the DsRed-TRF1 plasmid (Bronshtein et al., 2015) one day before imaging. For imaging, cells were placed in an incubator (Tokai, Shizuoka-Ken, Japan) mounted on an inverted Olympus IX-81 fluorescence microscope coupled to a FV-1000 confocal set-up (Olympus, Tokyo, Japan) using a UPLSAPO 60X objective lens with a numerical aperture of 1.35. Each nucleus was measured 50 times in three dimensions (3D) for a total time of 20.5 min. Imaris image analysis software package (Bitplane, Zurich, Switzerland) was used for correcting the nucleus drift and rotation and for identifying the coordinates of labeled telomeres. Only telomeres that were tracked over all the 50 time points were considered for further data analysis. For calculating the volume covered by each telomere during its whole motion, we used the Convex hull algorithm using a custom-made Matlab code.

Quantitative real-time PCR for detecting untagged and mEos3.2-tagged lamin A/C mRNA levels

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RNA was isolated from mEos3.2-lamin A/C WT#21 cells using the RNeasy mini plus kit (Qiagen) according to manufacturer’s instructions. cDNA was synthesized from 500 ng total RNA using the RevertAid First Strand cDNA synthesis kit (Thermo Fisher Scientific) and analyzed by qPCR using the KAPA SYBR Green 2x PCR master mix (Kapa Biosystems, Wilmington, MA) in an Eppendorf Realplex 2 Mastercycler according to manufacturer's instructions. Primers specific for untagged and tagged lamins A/C (wt LAC-f and -r; mEos LA/C-f and -r – see ‘Primers’) were used to generate PCR products that were gel-extracted and used as a template in real time PCR to generate a standard curve (DNA concentration versus threshold cycle). WT#21 cDNA was then analyzed by real-time PCR using the same primers and DNA/RNA concentration of the specific template was calculated from the standard curve.

Generation of Lmna-specific targeting construct

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The Lmna-specific targeting construct was assembled from four fragments that were amplified by PCR using the following primer pairs and templates (see also ‘Primers’):

FragmentF primer (5’- 3’)R primer (5’- 3’)Template
1mLmna-Frag1-fmLmna-Frag1-rMouse genomic DNA
2mEos-Frag2-fmEos-Frag2-rmEos3.2-C1 (Addgene, plasmid #54550)
3mLmna-Frag3-fmLmna-Frag3-rMouse genomic DNA
4mLmna-Frag4-fmLmna-Frag4-rMouse genomic DNA

Primers were designed to generate overlapping fragments that were assembled using the Gibson assembly master mix (New England Biolabs, Ipswich, MA) according to manufacturer’s instructions. The vector pUC18 (Norrander et al., 1983) was digested with SalI and EcoRI (New England Biolabs) creating vector ends overlapping with fragments 1 and 4, respectively, and added to the Gibson assembly reaction. The final Lmna targeting construct in pUC18 was sequence-verified and contained Lmna exon one and its flanking non-coding sequences, where mEos3.2 was inserted in frame into exon one before the first codon (removing the start codon). An additional EcoRI site was inserted directly after mEos3.2 and the recognition sequence of Lmna-specific sgRNA1 was altered within exon 1 (5’ CACTCGGATCACCCGcCTaC 3’, mutations are indicated in lowercase) to avoid recutting and potential creation of Indels in the modified allele. The Lmna targeting construct in pUC18 was amplified in high-efficiency NEB5-alpha competent E. coli (New England Biolabs).

Genotyping PCR and long range (LR) PCR for modified mEos3.2-Lmna knock-in allele

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To identify clones with a modified mEos3.2-Lmna knock-in allele, genomic DNA was isolated from single-cell clones using the QuickExtract DNA extraction solution (Epicentre/Lucigen, Middleton, WI) according to manufacturer’s instructions and analyzed for the presence of the wild-type and knock-in Lmna allele by PCR using the GoTaq green master mix (Promega, Madison, WI) and the following primer pairs (see also ‘Primers’):

PCRForward primer (5’- 3’)Reverse primer (5’- 3’)Product
WTmLmna-WT-fmLmna-WT-r213 bp (KI: 894 bp)
Knock-inmLmna-KI-fmLmna-KI-r250 bp

LR-PCR was performed in clones with at least one knock-in allele to verify correct integration of the construct at the 3-prime and 5-prime side with one primer of each pair outside the homology region of the Lmna targeting construct (see also Figure 2—figure supplement 1). PCR reactions were set up using the Q5 DNA polymerase (New England Biolabs) according to manufacturer’s instructions and the following primer pairs:

PCRForward primer (5’- 3’)Reverse primer (5’- 3’)Product
5-primeLmna-LR-5’-fLmna-LR-5’-r2566 bp
3-primeLmna-LR-3’-fLmna-LR-3’-r1710b p

Vectors

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pSpCas9(BB)−2A-mCherry was generated from pSpCas9(BB)−2A-GFP (Addgene, plasmid #48138) by removing EGFP and the T2A sequence via EcoRI digestion. The final vector was then assembled using the Gibson assembly master mix (New England Biolabs) and two overlapping fragments containing the T2A sequence and mCherry generated by PCR using the following primers and templates (see ‘Primers’):

FragmentForward primer (5’- 3’)Reverse primer (5’- 3’)Template
T2AT2A-fT2A-rpSpCas9(BB)−2A-GFP
mCherrymCherry-fmCherry-rpLVX mCherry (Takara Bio)

Vectors expressing different lamin A constructs were all derived from pEGFP-myc-LMNA (Moir et al., 2000). pEGFP-myc-Lamin A del K32, harboring a deletion of lysine at position 32 (Bertrand et al., 2012), was derived from pEGFP-myc-LMNA by site directed mutagenesis as previously described (Pilat et al., 2013). Wildtype and delK32 mature lamin A constructs were generated by PCR from pEGFP-myc-LMNA and pEGFP-myc-Lamin A delK32, respectively, using primers hLaminA-NheI-f and hLaminA-NheI-r.

Primers

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PrimerSequence (5’ - 3’)
hLAP2α-fACCTCAGGGCAACTTTAAAGCAA
hLAP2α-rTCTACATCCAGTGGGGGCATA
mLap2α-fGGGTCTTTTATGGGCCATTTTTGT
mLap2α-rCTCTTCCCTCCACGGCAAA
mLmna-fACCCCCTCCCTTCTATGTCC
mLmna-rGGAAGTGGGGTGAGTCACTG
wt LAC-fCTGCCGGCCATGGAGAC
wt LAC-rGCTGACCACCTCTTCAGACT
mEos LA/C-fCATGCTGTTGCTCATTCTGGA
mEos LA/C-rACAAGTCCCCCTCCTTCTTGG
mLmna-Frag1-fTTGCATGCCTGCAGGTCGACAATCTCTTAAGAGTCCCAACTCAAGCCCA
mLmna-Frag1-rTAATCGCACTCATGGCCGGCAGGGTGACAGT
mEos-Frag2-fGCCATGAGTGCGATTAAGCCAGACATGA
mEos-Frag2-rGGTCTCGAATTCTCGTCTGGCATTGTCAGGCAAT
mLmna-Frag3-fACGAGAATTCGAGACCCCGTCACAGCGG
mLmna-Frag3-rGTAGGCGGGTGATCCGAGTGGGCGA
mLmna-Frag4-fGGATCACCCGCCTACAAGAGAAGGAGGACCTGCAGGAGCT
mLmna-Frag4-rCTATGACCATGATTACGAATTCCCTAGGGCTGGAATCTGGTAAGGAA
mLmna-WT-fGATCGATGTACACGGCCAGG
mLmna-WT-rGTCCTTCTGTCCAAGTCCCG
mLmna-KI-fGATCGATGTACACGGCCAGG
mLmna-KI-rGTGGACCACTGCATTGAGATTTT
Lmna-LR-5’-fCCTAGGTTCCCTCCCCTAGAT
Lmna-LR-5’-rGGTAACTTGACACCCTTCTCCTTAG
Lmna-LR-3’-fGAGCATGCTGTTGCTCATTCT
Lmna-LR-3’-rCCCATCTGTGCACATGACCT
T2A-fGGCAAAAAAGAAAAAGGAATTCGGCAGTGGAGAGGGCAGA
T2A-rTCACTGGGCCAGGATTCTCCTCGA
mCherry-fAATCCTGGCCCAGTGAGCAAGGGCGAGGAGGAT
mCherry-rAGCGAGCTCTAGTTAGAATTCCTTGTACAGCTCGTCCATGCCGCCGGT
hLaminA-NheI-fGCGGCGGCTAGCATGGTGAGCAAGGGCGAGGAGC
hLaminA-NheI-rGTATATACTAGTAGGAGCGGGTGACCAG

Statistical analysis

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The effect size was estimated based on preliminary data or similar experiments performed in a different cell system and the software G*power (http://www.gpower.hhu.de/) was used to estimate the required sample size to achieve a minimum power of 0.8 (or higher, if experimentally feasible). Especially for predicted small effect sizes, large (>100) experimental sample sizes were chosen. Experiments were performed at least in triplicates. For statistical analysis, biological replicates (e.g. different clones of the same genotype) from all experiments were used and are stated in each figure legend as the number of n. If technical replicates (e.g. qPCR triplicates from the same sample) are displayed, it is stated in the figure legend. To summarize experimental data sets, average, standard deviation and standard error of the mean were calculated and displayed in bar graphs. In specific cases, box plots were chosen to display data to better visualize data distributions. In box plots the median was depicted within the first and third quartiles with the whiskers representing minimal and maximal datapoints excluding statistical outliers (according to the method of Tukey using the inter-quartile range). For statistical analysis, normal distribution of data was tested using the D’Agostino-Pearson and Shapiro Wilk normality test. Additionally, quantile-quantile plots were created and visually inspected for normal distribution. For normally distributed data sets, the two-tailed student’s t test was used for statistical analysis (unpaired or paired, if data points were matched in pairs). The F test was used to determine whether variances of data sets are equal. Data expressed as proportions are not normally distributed and were transformed using the arcsin transformation (transformed value = acrsin value). Ratios were transformed using the logarithmic transformation. If multiple comparisons were necessary (e.g. more than 2 data sets to compare with each other), ANOVA was used, including post-hoc tests for pairwise comparisons (Tukey). For non-normally distributed data, the Mann-Whitney U test was used, or, for multiple comparisons, the Kruskal-Wallis test.

Appendix 1

Online supplemental material

Figure 1—figure supplement 1 shows the characterization of HeLa LAP2α knockout clones created by CRISPR-Cas9 used in Figure 1. Figure 1—videos 14 show live-cell imaging of HeLa wildtype and LAP2α knockout cells expressing either GFP-pre-lamin A (Figure 1—videos 1 and 2, corresponding to Figure 1A, panel 1 and 2, and Figure 1—video 3 displaying an identically treated, second LAP2α knockout clone) or GFP-ΔK32 pre-lamin A (Figure 1—video 4, corresponding to Figure 1A, panel 3), imaged 5 hr post transfection. Figure 1—videos 58 show live-cell imaging of HeLa wildtype and LAP2α knockout cells expressing either GFP-pre-lamin A (Figure 1—videos 5 and 6, corresponding to Figure 1B, panel 1 and 2, and Figure 1—video 7 displaying an identically treated, second LAP2α knockout clone) or GFP-ΔK32 pre-lamin A (Figure 1—video 8, corresponding to Figure 1B, panel 3), imaged 24 hr post transfection. Figure 1—figure supplement 2 shows live cell imaging of HeLa cells expressing mature GFP-lamin A or mature GFP-ΔK32 lamin A (associated with Figure 1). Figure 1—videos 910 correspond to Figure 1—figure supplement 2A and Figure 1—videos 1112 correspond to Figure 1—figure supplement 2B. Figure 2—figure supplement 1 shows the characterization of mEos3.2-lamin A/C mouse dermal fibroblasts used in Figures 26. Figure 2—figure supplement 2 shows the characterization of isogenic mEos3.2-lamin A/C LAP2α knockout cell lines used in Figures 26. Figure 2—source data 1 contains raw data for single cell measurements of DAPI and lamin A/C nucleoplasmic/peripheral ratio shown in Figure 2G–H. Figure 2—figure supplement 3 compares the nucleoplasmic/peripheral signal ratios of lamin A/C and lamin B1 (associated with Figure 2G–H). Figure 3—figure supplement 1 shows immunofluorescence images of various LAP2α knockout and wildtype mEos3.2 cells with different antibodies. Figure 4—source data 1 contains raw data for constant photobleaching measurements of intranuclear lamin A/C depicted in Figure 4A. Figure 4—source data 2 contains raw data for fluorescence correlation spectroscopy (FCS) measurements of intranuclear lamin A/C shown in Figure 4B. Figure 4—figure supplement 1 shows lamin A/C extraction and lamin phosphorylation in wildtype and LAP2α knockout HeLa cells (associated with Figures 4 and 5). Figure 6—figure supplement 1 shows characterization of Lmna knockout and Lmna/Lap2α double knockout mouse dermal fibroblasts used in Figure 6. Figure 6—source data 1 contains raw data for measurements of telomere motion volume depicted in Figure 6.

Data availability

All data generated or analysed during this study are included in the manuscript and supporting files. Source data files have been provided for Figures 2G-H, Figure 4A-B and Figure 6A.

References

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    Lamina-associated polypeptide 2alpha binds intranuclear A-type lamins
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Decision letter

  1. Megan C King
    Reviewing Editor; Yale School of Medicine, United States
  2. Suzanne R Pfeffer
    Senior Editor; Stanford University School of Medicine, United States
  3. Megan C King
    Reviewer; Yale School of Medicine, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This manuscript describes new tools to investigate the attributes specific to nucleoplasmic versus lamina-integrated A-type lamins. Based on their findings, the authors develop a new model in which LAP2α influences the conformational state of A-type lamins, antagonizing stable lamin A filament assembly with ramifications for chromatin mobility.

Decision letter after peer review:

Thank you for submitting your article "LAP2α maintains a mobile and low assembly state of A-type lamins in the nuclear interior" for consideration by eLife. Your article has been reviewed by three peer reviewers, including Megan C King as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Suzanne Pfeffer as the Senior Editor.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). Specifically, when editors judge that a submitted work as a whole belongs in eLife but that some conclusions require a modest amount of additional new data, as they do with your paper, we are asking that the manuscript be revised to either limit claims to those supported by data in hand, or to explicitly state that the relevant conclusions require additional supporting data.

Our expectation is that the authors will eventually carry out the additional experiments and report on how they affect the relevant conclusions either in a preprint on bioRxiv or medRxiv, or if appropriate, as a Research Advance in eLife, either of which would be linked to the original paper.

Summary:

This work builds on prior studies by the Foisner group that investigated the function(s) of the soluble A-type lamin binding protein, LAP2α. One of their prior observations using antibody labeling was that there appeared to be a depletion of the nucleoplasmic pool of A-type lamins in cells lacking LAP2α. In this manuscript, the authors employ CRISPR-Cas9 editing to develop new tools to investigate the attributes specific to nucleoplasmic versus lamina-integrated A-type lamins. Using this new approach (and comparing it with their prior observations), the authors hit upon a new model in which LAP2α influences the conformational state of A-type lamins, which in turn influences its detection by a commonly used antibody. This technical detail explains the new realization that nucleoplasmic lamin A persists in LAP2α-null cells, albeit in a different state. The authors provide evidence that LAP2α antagonizes stable lamin A filament assembly, that is absence leads to stabilized intranuclear lamin A assemblies, and that telomere mobility is negatively influenced by loss of LAP2α in a manner depending on the presence of lamin A/C. The authors' work further identifies two pathways by which nucleoplasmic lamins emerge, namely by 1) initial localization to the lamina followed by relocalization to the nucleoplasm, and 2) from the pool of mitotic lamins which are not associated to the lamina.

Overall there was enthusiasm for the study, with the reviewers stating their appreciation for the author's mechanistic approach to studying lamin assembly state and the use of complementary cell biology/microscopy and biochemical approaches. The rigor of the science was also lauded, including inclusion of, for example, genome editing quality control measures. Taken together the reviewers felt that the findings provided a new perspective on how LAP2α influences the state of A-type lamins. As the impact of lamins on nuclear organization is critical for nuclear functions and important for nuclear integrity, these results are fundamental for the understanding of both lamin A/C and LAP2α. The reviewers also made specific suggestions for a small number of experimental revisions as well as further editing of the text to improve clarity and to provide greater context.

Essential revisions:

1) Structure of the Results. Given that a major focus of the paper is to explain conflicting results with (the same group's) prior published data on the effect of LAP2α depletion, it would have helped to lay this out more clearly from the outset of the paper. As written, the reader is confused until arriving at Figure 3. While the reviewers appreciate that resolving this conflict leads to a new perspective – namely that LAP2α influences the state of the lamin assembly in a way that disrupts its detection by the N18 antibody, structuring the manuscript to get to this point as quickly as possible would improve its accessibility.

2) Suggested edits to clarify the manuscript. Although the reviewers appreciated the transparency of the authors in demonstrating their workflow and quality control measures, some of the terminology made the manuscript difficult to read. For example, the manuscript would be easier to understand if cell lines were given descriptive names (e.g.: LAP2α KO, or mEos3.2-lmna instead of "WT#21") rather than continuing to refer to them by the small guide RNA that was used to generate them. A second example: while presenting biological replicate data as in Figure 1 is appreciated, it was challenging to ascertain that the second and third columns in panels A and B were biological replicates. The authors could improve clarity by presenting one biological replicate in the main text with additional replicate(s) moved to the supplement, especially considering that it appears that only one of the clones was used for the quantifications shown in the bottom panels.

3) Alternative explanations. Further comment and/or clarification is necessary when discussing the state of nucleoplasmic lamin polymerization and how the dynamic measurements reflect lamin assembly state versus anchorage to other lamin binding proteins. For example, although the nuclear lamina filaments are typically 200-400 nm in length, they are also very flexible. A 200 nm filament would have a molecular weight of <1.4MDa ( ~50% of a ribosome) and can be bent and curved. As a consequence, a single filament would be expected to have a reasonably high diffusion coefficient. At the lamina, lamins are less mobile, however, it is likely to be due to binding partners that anchor lamins to the INM and chromatin rather than assembly state as the diffusion of even a 1.4 MDa protein complexes is quite fast. Thus, the authors need to be careful in their interpretation as nucleoplasmic lamins may be polymerized but more mobile (less anchored) than their lamina-bound lamin population without a change in assembly state. Along these lines, greater explanation for the apparent paradox between the increase in immobile fraction by FRAP and the increased diffusion coefficient by FCS in the LAP2α-depleted condition is needed. The authors suggest that the latter is due to the loss of LAP2α binding (subsection “LAP2α loss changes properties of nucleoplasmic lamin A/C rendering them less mobile and more resistant to biochemical extraction”), but some modeling would go a long way here. What form are the lamins thought to be in, and how does the bulk that LAP2α would bring match the apparent changes in diffusivity?

4) Edited clones. It was noted that the authors chose to disrupt LAP2α at the beginning of exon 4, but the reason for this was unclear. For the readers' clarity, can the authors include the size of exons 1 and 2 in the schematic in the supplementary figures? Can the authors provide evidence that the "LAP2α KO" cells used in the experiments do not express a truncated LAP2α protein that could influence the results and interpretation?

5) Concerns over whether lamin tagging may influence the results. The authors make no comment on the functionality of the mEos-tagged lamin A/C CRISPR lines. However, the comment suggesting that some clones could have altered nuclear morphology (subsection “LAP2α-deficiency does neither affect the formation nor the steady state level of endogenous lamin A in the nuclear interior”) raises some questions. How did the authors interpret this? Were these clones in which there were indels in some lmna alleles affecting the levels? Or is this a consequence of the fusion? One approach we suggest is to compare morphology and overall fitness of cells with all untagged lmna with cells with all tagged lmna, to determine whether the tagged proteins are fully functional. How do the authors explain the relatively low expression level of the mEos fusion relative to the untagged? If the MDFs are diploid, presumably we would expect this to be one allele tagged and one allele untagged. Given that the expression ratio is very different from this, could the tagged lamin A/C be targeted for degradation? As these cell lines are critical for the rest of the study, this information is important, particularly as many lamin mutations do not cause obvious phenotypes in tissue culture cells, but defects can still emerge during development and aging in the context of an animal. Further, in the subsection “LAP2α loss changes properties of nucleoplasmic lamin A/C rendering them less mobile and more resistant to biochemical extraction”, the authors describe a set of experiments that are meant to demonstrate that their failure to see a difference in nucleoplasmic A-type lamins in LAP2α mutants is not due to the fluorescent protein tag used, however, instead of looking at untagged lamins, they elect to look at a cell line that has all lmna alleles tagged. The authors' statements would be strengthened if the LAP2α KO cells from Figure 1 were stained with both the 3A6 antibody and the N18 antibody to determine whether untagged lamins behave the same way as tagged lamins. More generally, including experiments that allow the reader to compare directly between a cell line with all lmna alleles tagged and a cell line with no lmna alleles tagged is important.

6) Implications for regulation of A-type lamin assembly. The authors build a convincing case that binding to A-type lamins by LAP2α influences their ability to assemble. But how do cells leverage this relationship for biological functions? Do cells tune the amount of fully soluble vs. partially assembled A-type lamins in the nucleoplasm in order to control nuclear structure or function in response to certain stimuli (e.g. cyclical stretch, for example)? Have the A-type lamins in the nucleoplasm been found to be in a different assembly state in different cell types? Similarly, one prediction that arises from the proposed model is that regulation of LAP2α levels will modulate the relative pool of A-type lamins at the nuclear interior versus the nucleoplasm. Beyond the knock-out cells, is there any other evidence of this relationship? How do the authors think the membrane integrated LAP2β fits into the story? Any further insight provided would greatly help to address biological mechanism.

7) Lamin assembly mechanism. The authors show that nucleoplasmic lamins are first localized to the lamina, where they can polymerize. Isn't it possible that filaments can be released into the nucleoplasm? The authors suggest that Lap2α keeps lamin in a less polymerized state based in part on the findings that Lap2α inhibits lamin A assembly in vitro. However, previous work by Zwerger et al., 2015, showed that inhibitors of in vitro lamin A assembly have no impact on incorporation and localization of lamin A into the lamina, while incorporation of lamin A into the nuclear lamina was abolished when other lamin binders that have no effect on lamin assembly in vitro were used. This suggests that either in vitro assembly is not representing the cellular lamin assembly or assembly of lamin into the lamina is independent of polymerization states of lamins. The authors should discuss these views in the revised manuscript. Greater use of the ΔK32 mutant was also suggested, specifically as related to Figure 1, where the data on this assembly mutant should be included in the primary figures and with regards to the sensitivity of the N18 antibody – does this detect the ΔK32 mutated lamin A? Could this provide further insight into the impact of LAP2α by extension?

8) Chromatin dynamics. Over what time interval was the volume of movement for the telomeres observed? This is important because more fluctuations in nuclear position, for example, could influence this measure. In addition, telomeres are a confusing choice, given abundant evidence that there is crosstalk between the state of the nuclear lamina and telomere biology (e.g. lamin mutants affecting telomere homeostasis, etc.). At a minimum, acknowledging that telomeres may not reflect the effect on chromatin globally is important. Examples of the raw mean squared displacements would be more informative. Is the difference between lmna KO and lmna/Lap2α DKO (Figure 6 right panel) significant?

https://doi.org/10.7554/eLife.63476.sa1

Author response

Essential revisions:

1) Structure of the Results. Given that a major focus of the paper is to explain conflicting results with (the same group's) prior published data on the effect of LAP2α depletion, it would have helped to lay this out more clearly from the outset of the paper. As written, the reader is confused until arriving at Figure 3. While the reviewers appreciate that resolving this conflict leads to a new perspective – namely that LAP2α influences the state of the lamin assembly in a way that disrupts its detection by the N18 antibody, structuring the manuscript to get to this point as quickly as possible would improve its accessibility.

We understand the concerns of the reviewers and have now rewritten the Abstract and the last paragraph in the Introduction to more clearly point out the aims of this study and the surprising finding that led to a new adapted model of LAP2α function in the regulation of nucleoplasmic lamins. In addition, we streamlined the text describing the formation of the nucleoplasmic lamin pool in the Results (Figures 1 and 2) and changed the focus more towards the effect of LAP2α rather than the pathways involved in the formation of lamins A/C in the nuclear interior.

We also considered rearranging the figures, but after extensive discussion we prefer to keep the current order of figures, as rearrangements would make the structure of the manuscript less clear and disturb the logical flow of the text and experimental set up (e.g. ectopic proteins versus endogenously labeled lamin A/C). We are confident that with the included text changes we can best address the reviewer’s comments, while maintaining the order of the figures.

2) Suggested edits to clarify the manuscript. Although the reviewers appreciated the transparency of the authors in demonstrating their workflow and quality control measures, some of the terminology made the manuscript difficult to read. For example, the manuscript would be easier to understand if cell lines were given descriptive names (e.g.: LAP2α KO, or mEos3.2-lmna instead of "WT#21") rather than continuing to refer to them by the small guide RNA that was used to generate them. A second example: while presenting biological replicate data as in Figure 1 is appreciated, it was challenging to ascertain that the second and third columns in panels A and B were biological replicates. The authors could improve clarity by presenting one biological replicate in the main text with additional replicate(s) moved to the supplement, especially considering that it appears that only one of the clones was used for the quantifications shown in the bottom panels.

We now use descriptive names for the cell lines throughout the text and in most figures. In figures, where different LAP2α knockout clones generated by different sgRNAs were used, as well as cell clones treated with the same sgRNAs, but still expressing LAP2α, we designated the cells LAP2α KO sgxx and WT sgxx ctrl (see for example Figure 2F and G). These designations are clearly explained in the figure legend of Figure 2. We agree that these changes make it much easier for a broad readership to follow the logical flow of the manuscript.

In order to address the point of the biological replicate, we removed replicate panels in Figure 1A and B. Instead of showing replicate panels in Figure 1, we now include these data in additional video files (Videos 3 and 7).

3) Alternative explanations. Further comment and/or clarification is necessary when discussing the state of nucleoplasmic lamin polymerization and how the dynamic measurements reflect lamin assembly state versus anchorage to other lamin binding proteins. For example, although the nuclear lamina filaments are typically 200-400 nm in length, they are also very flexible. A 200 nm filament would have a molecular weight of <1.4MDa ( ~50% of a ribosome) and can be bent and curved. As a consequence, a single filament would be expected to have a reasonably high diffusion coefficient. At the lamina, lamins are less mobile, however, it is likely to be due to binding partners that anchor lamins to the INM and chromatin rather than assembly state as the diffusion of even a 1.4 MDa protein complexes is quite fast. Thus, the authors need to be careful in their interpretation as nucleoplasmic lamins may be polymerized but more mobile (less anchored) than their lamina-bound lamin population without a change in assembly state. Along these lines, greater explanation for the apparent paradox between the increase in immobile fraction by FRAP and the increased diffusion coefficient by FCS in the LAP2α-depleted condition is needed. The authors suggest that the latter is due to the loss of LAP2α binding (subsection “LAP2α loss changes properties of nucleoplasmic lamin A/C rendering them less mobile and more resistant to biochemical extraction”), but some modeling would go a long way here. What form are the lamins thought to be in, and how does the bulk that LAP2α would bring match the apparent changes in diffusivity?

We toned down the conclusions on lamin polymerization and included alternative explanations based on lamin filament anchorage to explain the observed differences in lamin A mobility (i.e. movement) in LAP2α KO versus WT cells throughout the text and added a paragraph on this topic in the Discussion.

The increase in the immobile fraction in the absence of LAP2α as measured by constant photobleaching suggests that a larger fraction of intranuclear lamins are immobile, likely because they are bound to an entity that prevents its motion (e.g. chromatin or larger lamin filament assemblies in the nucleoplasm). These immobile structures cannot be measured by FCS, as their fluorescence intensity is not fluctuating. Hence, FCS data refer exclusively to the unbound, mobile fraction of lamin A, which is reduced (but still present) in LAP2α knockout cells. This mobile lamin A/C fraction may be diffusing faster as the complexes without LAP2α are smaller than in WT cells. In the manuscript we explain the results more clearly and give an estimation of a potential correlation of diffusion coefficient changes with changes in the molecular mass of the remaining dynamic lamin complex in the absence of LAP2α. The exact form these mobile lamin-LAP2α complexes are in is unknown. For the calculations, we are assuming the simplest form with one molecule each present in the complexes. Notably, our unpublished data suggest a stoichiometry of 1:1 or 1:2 for soluble lamin A/C-LAP2α complexes.

4) Edited clones. It was noted that the authors chose to disrupt LAP2α at the beginning of exon 4, but the reason for this was unclear. For the readers' clarity, can the authors include the size of exons 1 and 2 in the schematic in the supplementary figures? Can the authors provide evidence that the "LAP2α KO" cells used in the experiments do not express a truncated LAP2α protein that could influence the results and interpretation?

We now clarify in the figure legend that exon 4 is the first and only LAPα-specific exon in the LAP2 gene, while exons 1-3 are common to all LAP2 isoforms. Thus, generating a LAP2α-specific knockout without disturbing expression of the other isoforms requires targeting the beginning of exon 4. To address the reviewer’s specific point, we also added exons 1 to 3 in the schematic drawings of Figure 1—figure supplement 1 and Figure 2—figure supplement 2.

As for a truncated LAP2α fragment, one LAP2α KO clone (sg3) indeed expresses low levels of a small LAP2α fragment (see Figure 2—figure supplement 2D), but we mostly used sg1 clones for the analyses, which do not contain a fragment (Figure 2—figure supplement 2D). In experiments, where sg3 was used (data in Figure 2E-G, Figure 3—figure supplement 1A), the respective data are clearly labeled and explained in the legend, and we included a heterozygous clone that was treated with sg3 (former clone sg3-2, now labeled with WT sg3 ctrl) as control, expressing LAP2α from 1 allele and a short truncated LAP2α fragment from another targeted allele (see Figure 2—figure supplement 2C and D). WT sg3 ctrl expresses full length LAP2α at WT levels (Figure 2B).

5) Concerns over whether lamin tagging may influence the results. The authors make no comment on the functionality of the mEos-tagged lamin A/C CRISPR lines. However, the comment suggesting that some clones could have altered nuclear morphology (subsection “LAP2α-deficiency does neither affect the formation nor the steady state level of endogenous lamin A in the nuclear interior”) raises some questions. How did the authors interpret this? Were these clones in which there were indels in some lmnA alleles affecting the levels? Or is this a consequence of the fusion? One approach we suggest is to compare morphology and overall fitness of cells with all untagged lmna with cells with all tagged lmna, to determine whether the tagged proteins are fully functional. How do the authors explain the relatively low expression level of the mEos fusion relative to the untagged? If the MDFs are diploid, presumably we would expect this to be one allele tagged and one allele untagged. Given that the expression ratio is very different from this, could the tagged lamin A/C be targeted for degradation? As these cell lines are critical for the rest of the study, this information is important, particularly as many lamin mutations do not cause obvious phenotypes in tissue culture cells, but defects can still emerge during development and aging in the context of an animal. Further, in the subsection “LAP2α loss changes properties of nucleoplasmic lamin A/C rendering them less mobile and more resistant to biochemical extraction”, the authors describe a set of experiments that are meant to demonstrate that their failure to see a difference in nucleoplasmic A-type lamins in LAP2α mutants is not due to the fluorescent protein tag used, however, instead of looking at untagged lamins, they elect to look at a cell line that has all lmna alleles tagged. The authors' statements would be strengthened if the LAP2α KO cells from Figure 1 were stained with both the 3A6 antibody and the N18 antibody to determine whether untagged lamins behave the same way as tagged lamins. More generally, including experiments that allow the reader to compare directly between a cell line with all lmna alleles tagged and a cell line with no lmna alleles tagged is important.

The clone used in most experiments (mEos3.2-Lmna WT clone #21, see Figure 2—figure supplement 1D and F) is tetraploid (as determined from sequencing data and TIDE analysis, see Figure 2—figure supplement 2B and C) and expresses tagged lamin A/C from 1 allele and untagged lamin A/C from 3 alleles (see Figure 2—figure supplement 1G for ratio of tagged versus untagged lamin A mRNA and protein levels in newly added protein quantification).

There may be a certain influence of the tag on protein stability, as the ratio of tagged to untagged lamin A/C protein is shifted towards approx. 1:8 on the protein level (Figure 2—figure supplement 1G), but functions and properties of tagged lamin A and C seem to be unaffected. In the manuscript, we refer to the correct localization of tagged lamin A (Figure 2C), to the stable integration into the lamina by FRAP (former Figure 2—figure supplement 1H, now moved to Figure 2 as new panel D), normal post-mitotic lamina assembly (shown in Figure 2) and similar biochemical properties, such as solubility in salt/detergent buffer (newly added quantification of solubility in Figure 2—figure supplement 1G, right panel) (subsection “LAP2α-deficiency does neither affect the formation nor the steady state level of endogenous lamin A in the nuclear interior”).

However, while some mEos3.2-Lmna clones (independent of whether WT lamin A/C was still present or not) had altered nuclear morphology, this often correlated with very low total lamin A/C levels due to indels in some Lmna alleles as also pointed out by the reviewers. We chose two clones expressing only mEos-tagged lamin A/C, one WT and one KO for LAP2α (WT#5 and LAP2α KO#17, see Figure 2—figure supplement 1D and F) for further control experiments, as suggested by the reviewers. The WT clone expressing only tagged lamin A/C has lower total levels of lamin A/C (likely expressed only from 1 allele while other alleles contain indels), but still displays normal nuclear morphology (see Figure 2—figure supplement 1I). Following the suggestion of the reviewers to test the fitness of these cell clones and to confirm that tagged lamin A/C behaves like untagged lamins, we added additional data comparing the parental WT and LAP2α KO cells (no tagged lamins) with the cell clones expressing only tagged lamin A/C regarding cell viability, nuclear morphology, lamin localization and lamina structure (Figure 2—figure supplement 1H and I). Notably, WT#5 cells expressing lower total levels of mEos-tagged lamins A/C displays a slight reduction in cell viability (pointed out in the figure legend), which is likely caused by reduced total lamin levels rather than the tagging of lamins, as LAP2α KO#17 cells expressing higher mEos-lamin A/C levels display normal viability, when compared to parental cells with untagged Lmna alleles (Figure 2—figure supplement 1H).

Additionally, we performed immunofluorescence microscopic analyses of these cells using anti-lamin A/C N18 and 3A6 antibody and confirmed the lack of intranuclear lamin staining in the absence of LAP2α in both, cells without tagged lamins and cell clones expressing only tagged lamins (Figure 3—figure supplement 1C and D). Thus, by all these means cells with only tagged lamins behave as cells with untagged lamins.

We also discuss functionality of tagged lamin A/C now in more detail in the Results section.

6) Implications for regulation of A-type lamin assembly. The authors build a convincing case that binding to A-type lamins by LAP2α influences their ability to assemble. But how do cells leverage this relationship for biological functions? Do cells tune the amount of fully soluble vs. partially assembled A-type lamins in the nucleoplasm in order to control nuclear structure or function in response to certain stimuli (e.g. cyclical stretch, for example)? Have the A-type lamins in the nucleoplasm been found to be in a different assembly state in different cell types? Similarly, one prediction that arises from the proposed model is that regulation of LAP2α levels will modulate the relative pool of A-type lamins at the nuclear interior versus the nucleoplasm. Beyond the knock-out cells, is there any other evidence of this relationship? How do the authors think the membrane integrated LAP2β fits into the story? Any further insight provided would greatly help to address biological mechanism.

In the revised manuscript, we discuss a potential biological function in great detail.

As we have shown previously, LAP2α loss clearly impairs the detection of nucleoplasmic lamins by antibodies in various cell types, including fibroblasts and epidermal and colon progenitor cells (Naetar et al., 2008). Changes in antibody-stained lamin A/C levels in the nuclear interior can also be seen in physiological conditions within cell populations in a cell culture, expressing different levels of LAP2α (see e.g. Figure 1—figure supplement 1E, depicting cells with different levels of LAP2α side by side).

As for a potential biological function and regulation of nucleoplasmic lamins we discuss the following points: While the response of lamin A solubility to mechanical stretch works via lamin A phosphorylation (Buxboim et al., 2014), changes in the detectability of lamin A/C in the nuclear interior during differentiation may be regulated by differentiation-specific downregulation of LAP2α (Markiewicz, Ledran, and Hutchison, 2005; Naetar et al., 2007; Pekovic et al., 2007). Similar effects can be seen in progeria premature aging disease (Vidak et al., 2018; Vidak et al., 2015). We hypothesize that a mobile nucleoplasmic lamin A is important for chromatin regulation and gene expression [see (Bronshtein et al., 2015; Gesson et al., 2016), our unpublished data, and shown in this manuscript in Figure 6], as well as for the regulation of retinoblastoma protein (Dorner et al., 2006) during cell cycle exit and initiation of cell differentiation.

LAP2β was shown to bind lamin B, not lamin A/C (Foisner and Gerace, 1993), while LAP2α binds lamin A/C via a LAP2α-specific domain encoded by exon 4 (Dechat et al., 2000) absent in LAP2β. Thus, we feel that the function of LAP2α regulating lamin A/C in the nuclear interior is unique to the protein and not affected by the other LAP2 isoforms.

7) Lamin assembly mechanism. The authors show that nucleoplasmic lamins are first localized to the lamina, where they can polymerize. Isn't it possible that filaments can be released into the nucleoplasm? The authors suggest that Lap2α keeps lamin in a less polymerized state based in part on the findings that Lap2α inhibits lamin A assembly in vitro. However, previous work by Zwerger et al., 2015, showed that inhibitors of in vitro lamin A assembly have no impact on incorporation and localization of lamin A into the lamina, while incorporation of lamin A into the nuclear lamina was abolished when other lamin binders that have no effect on lamin assembly in vitro were used. This suggests that either in vitro assembly is not representing the cellular lamin assembly or assembly of lamin into the lamina is independent of polymerization states of lamins. The authors should discuss these views in the revised manuscript. Greater use of the ΔK32 mutant was also suggested, specifically as related to Figure 1, where the data on this assembly mutant should be included in the primary figures and with regards to the sensitivity of the N18 antibody – does this detect the ΔK32 mutated lamin A? Could this provide further insight into the impact of LAP2α by extension?

We discuss now in great detail that lamin A assembly may be different in vivo versus in vitro, in view of the data shown in Zwerger et al., 2015, and we also describe alternative models on the assembly state and interactions of nucleoplasmic lamins (see also our response to point 3 above) (Discussion).

In order to highlight the results of the assembly-deficient ΔK32 prelamin A mutant we moved panels showing assembly of this lamin mutant from Figure 1—figure supplement 2 to Figure 1.

Stainings of ΔK32 lamin A mutant by the anti-lamin A/C N18 antibody in LAP2α WT versus KO cells, as mentioned by the reviewer, were already done and shown in one of our previous papers (Pilat et al., 2013). As expected, unlike WT lamin A, the intranuclear staining of the assembly-deficient ΔK32 lamin A by antibody N18 was not impaired in LAP2α KO cells, which would be in support of lamin A assembly being responsible for loss of N18 staining. We add these findings in the Discussion.

8) Chromatin dynamics. Over what time interval was the volume of movement for the telomeres observed? This is important because more fluctuations in nuclear position, for example, could influence this measure. In addition, telomeres are a confusing choice, given abundant evidence that there is crosstalk between the state of the nuclear lamina and telomere biology (e.g. lamin mutants affecting telomere homeostasis, etc.). At a minimum, acknowledging that telomeres may not reflect the effect on chromatin globally is important. Examples of the raw mean squared displacements would be more informative. Is the difference between lmna KO and lmna/Lap2α DKO (Figure 6 right panel) significant?

Each nucleus is measured for 20.5 minutes. Our extensive experience on measurements in this time range has shown that this is long enough to prevent that measurements are not affected by short-time motion, which may not represent the actual dynamics. [see also (Zada et al., 2019)].

Furthermore, the motion of the nucleus itself (“nuclear position”) is completely controlled during the analysis by performing a drift-correction algorithm to each measured cell. We use a robust algorithm (see for example: (Bronshtein et al., 2015), section methods; and in “Single particle tracking for studying the dynamic properties of genomic regions in live cells“ , I. Bronshtein Berger, E. Kepten and Y. Garini, In: Imaging gene expression Methods and Protocols, edited by Y. Shav-Tal, Springer 2013). Due to the fact that we can track simultaneously (over 20.5 minutes) at least 20-40 telomeres in each cell, and because they are spread in the nucleus volume, the drift-correction algorithm leads to excellent correction of the cell motion. This is done by calculating for each image (at each time point) the “center of gravity” of the nucleus and shifting the coordinates of the nucleus to the same very point. Only then we calculate the dynamics of each telomere, and therefore, the cell motion is not affecting the calculated track of the telomeres and the calculations of the MSD. In addition, we also perform a rotation-correction of each nucleus by using an adequate algorithm that is part of the routine analysis. These two corrections ensure that the MSD that we calculate is accurate and it is not affected by cell drift or rotation.

Regarding the choice of telomeres, we have performed numerous measurements of the dynamics of telomeres in different cell types (MEFs, U2OS and HeLa) while depleting different proteins, and both telomeres and centromeres in WT and lamin A-depleted U20S cells. It is our experience that the dynamics of both types of probes in all these cells behave in a similar manner, i.e. if the telomere MSD reduces, also the centromere MSD reduces and vice versa (Bronshtein et al., 2015). Therefore, we strongly believe that the effect that we measure is not related to the biological properties of the telomeres themselves, and they only serve as ‘beacons’ for observing the general dynamics of chromatin in the nucleus. Possible modifications to the telomeres biology itself that may result from protein binding or other modifications, may change the dynamics of the telomeres on the scale of sub-telomere size, most probably in a 10-20 nm motion range, while the volume of motion that we measure is roughly a sphere with a diameter of ~250-300 nm. Local internal-dynamics of the telomere cannot affect the volume of its motion as a whole, which is what we are measuring.

As requested, for clarifying the data to the readers, we are adding a graph that shows the MSD of telomeres in WT and LAP2α knockout cells to Figure 6 (new panel B). The graph clearly shows that the absence of LAP2α leads to a ‘slower’, more constraint diffusion with lower MSD. These data are correlated with the change in the volume of motion shown in Figure 6A.

We also calculated the corrected p value for comparing Lmna KO and Lmna/Lap2α DKO, which is highly significant (Dunn's multiple comparisons test p < 0.0001 as part of Kruskal-Wallis post tests). Despite the rather small effect size (Cohen’s d=0.3), this suggests that LAP2α has a small, but significant effect on chromatin motion independent of lamin A/C, where its presence further restricts free diffusion of chromatin. This is indicated by the increase in chromatin motion volume in DKO compared to Lmna single KO cells (Figure 6A). These observations were included in the Results and Discussion sections of the manuscript.

References:

Foisner, R., and Gerace, L. (1993). Integral membrane proteins of the nuclear envelope interact with lamins and chromosomes, and binding is modulated by mitotic phosphorylation. Cell, 73(7), 1267-1279. doi:10.1016/0092-8674(93)90355-tPekovic, V., Harborth, J., Broers, J. L., Ramaekers, F. C., van Engelen, B., Lammens, M., von Zglinicki, T., Foisner, R., Hutchison, C., and Markiewicz, E. (2007). Nucleoplasmic LAP2alpha-lamin A complexes are required to maintain a proliferative state in human fibroblasts. J Cell Biol, 176(2), 163-172. doi:10.1083/jcb.200606139Vidak, S., Georgiou, K., Fichtinger, P., Naetar, N., Dechat, T., and Foisner, R. (2018). Nucleoplasmic lamins define growth-regulating functions of lamina-associated polypeptide 2alpha in progeria cells. J Cell Sci, 131(3). doi:10.1242/jcs.208462Zada, D., Bronshtein, I., Lerer-Goldshtein, T., Garini, Y., and Appelbaum, L. (2019). Sleep increases chromosome dynamics to enable reduction of accumulating DNA damage in single neurons. Nat Commun, 10(1), 895. doi:10.1038/s41467-019-08806-w

https://doi.org/10.7554/eLife.63476.sa2

Article and author information

Author details

  1. Nana Naetar

    Max Perutz Labs, Center for Medical Biochemistry, Medical University of Vienna, Vienna Biocenter Campus (VBC), Vienna, Austria
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Writing - original draft, Writing - review and editing
    Contributed equally with
    Konstantina Georgiou
    For correspondence
    nana.naetar@univie.ac.at
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8978-466X
  2. Konstantina Georgiou

    Max Perutz Labs, Center for Medical Biochemistry, Medical University of Vienna, Vienna Biocenter Campus (VBC), Vienna, Austria
    Contribution
    Formal analysis, Investigation, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Nana Naetar
    Competing interests
    No competing interests declared
  3. Christian Knapp

    Max Perutz Labs, Center for Medical Biochemistry, Medical University of Vienna, Vienna Biocenter Campus (VBC), Vienna, Austria
    Present address
    ICFO – Institut de Ciencies Fotoniques, The Barcelona Institute of Science and Technology, Castelldefels, Spain
    Contribution
    Software, Formal analysis, Investigation, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2463-5775
  4. Irena Bronshtein

    Physics Department and Nanotechnology Institute, Bar Ilan University, Ramat Gan, Israel
    Contribution
    Formal analysis, Investigation, Methodology, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
  5. Elisabeth Zier

    Max Perutz Labs, Center for Medical Biochemistry, Medical University of Vienna, Vienna Biocenter Campus (VBC), Vienna, Austria
    Contribution
    Formal analysis, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  6. Petra Fichtinger

    Max Perutz Labs, Center for Medical Biochemistry, Medical University of Vienna, Vienna Biocenter Campus (VBC), Vienna, Austria
    Contribution
    Resources, Investigation
    Competing interests
    No competing interests declared
  7. Thomas Dechat

    Max Perutz Labs, Center for Medical Biochemistry, Medical University of Vienna, Vienna Biocenter Campus (VBC), Vienna, Austria
    Present address
    Ludwig Boltzmann Institute of Osteology at Hanusch Hospital of OEGK and AUVA Trauma Centre Meidling, 1(st) Medical Department Hanusch Hospital, Vienna, Austria
    Contribution
    Conceptualization, Supervision, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3236-7889
  8. Yuval Garini

    Physics Department and Nanotechnology Institute, Bar Ilan University, Ramat Gan, Israel
    Present address
    Biomedical Engineering Department, Technion - Israel Institute of Technology, Haifa, Israel
    Contribution
    Conceptualization, Resources, Supervision, Validation, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8783-2015
  9. Roland Foisner

    Max Perutz Labs, Center for Medical Biochemistry, Medical University of Vienna, Vienna Biocenter Campus (VBC), Vienna, Austria
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Validation, Project administration, Writing - review and editing
    For correspondence
    roland.foisner@meduniwien.ac.at
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4734-4647

Funding

Austrian Science Fund (P26492-B20)

  • Roland Foisner

Austrian Academy of Sciences (APART 11657)

  • Nana Naetar

Israel Science Foundation (ISF grant 1219/17)

  • Irena Bronshtein
  • Yuval Garini

European Cooperation in Science and Technology (COST-STSM-BM1002-8698)

  • Thomas Dechat

European Molecular Biology Organization (ASTF 316-2011)

  • Thomas Dechat

S Grosskopf Grant

  • Irena Bronshtein
  • Yuval Garini

Austrian Academy of Sciences (ÖAW DOC 25725)

  • Konstantina Georgiou

Austrian Science Fund (P29713-B28)

  • Roland Foisner

Austrian Science Fund (P32512-B)

  • Roland Foisner

European Cooperation in Science and Technology (COST-STSM-BM1002-11436)

  • Thomas Dechat

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We are grateful to Harald Herrmann, DKFZ Heidelberg, Germany for the generous gift of recombinant lamin A and for his advice for lamin A in vitro assembly assays, and to Egon Ogris, Max Perutz Labs, Vienna, for the lamin A 3A6 antibody. We thank the Max Perutz Labs Biooptics facility for technical support with microscopic imaging and image analysis and technical support with FACS sorting. This study was funded by the Austrian Science Fund (FWF grant P26492-B20, P29713-B28 and P32512-B) to RF. KG is a recipient of a DOC Fellowship of the Austrian Academy of Sciences at the Max Perutz Labs, Medical University Vienna (ÖAW DOC 25725). NN was a recipient of an APART Fellowship of the Austrian Academy of Sciences at the Max Perutz Labs, Medical University Vienna (APART 11657). YG and IB acknowledge financial support from the Israel Science Foundation (ISF) grant 1219/17 and from the S Grosskopf grant for ‘Generalized dynamic measurements in live cells’. TD was a recipient of two COST Short Term Scientific Mission Fellowships (COST-STSM-BM1002-8698 and COST-STSM-BM1002-11436) and an EMBO short term fellowship (ASTF 316–2011).

Senior Editor

  1. Suzanne R Pfeffer, Stanford University School of Medicine, United States

Reviewing Editor

  1. Megan C King, Yale School of Medicine, United States

Reviewer

  1. Megan C King, Yale School of Medicine, United States

Publication history

  1. Received: September 28, 2020
  2. Accepted: February 18, 2021
  3. Accepted Manuscript published: February 19, 2021 (version 1)
  4. Version of Record published: March 8, 2021 (version 2)

Copyright

© 2021, Naetar et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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