Abstract
Stress granules (SGs) are cytoplasmic, membraneless organelles that
modulate mRNA metabolism and cellular adaptation under stress, yet the
mechanisms by which they regulate cancer cell survival remain unclear.
Here, we identify Poly(A)-Binding Protein Cytoplasmic 1 (PABPC1), a
core SG component, as stress-inducible SUMOylation target. Upon various
stress conditions, SUMOylated PABPC1 promotes SG assembly and enhances
cancer cell survival. Transcriptome-wide analysis reveals that
SUMOylated PABPC1 selectively stabilizes mRNAs enriched in conserved
U-rich elements. Mechanistically, SUMOylated PABPC1 interacts with
RNA-binding protein TIA1 to form PABPC1–SUMO–TIA1 complex that recruits
U-rich mRNAs into SGs, protecting them from degradation. This process
facilitates the expression of U-rich genes, such as mitophagy-related
genes FUNDC1, BNIP3L, thereby maintaining cellular homeostasis and
promoting cell survival under adverse conditions. Our findings reveal
that PABPC1 SUMOylation connects stress granule assembly with selective
U-rich mRNA stabilization and mitophagy, promoting cancer cell stress
adaptation.
Subject terms: RNA decay, Mechanisms of disease, Stress signalling,
Sumoylation
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Poly(A)-Binding Protein Cytoplasmic 1 (PABPC1) is a crucial component
of stress granules. Here, the authors show that PABPC1 undergoes
SUMOylation in response to cellular stress, enhancing the stability of
mitophagy-related gene transcripts to promote cancer cell survival.
Introduction
Eukaryotic cells encounter various stressors, including oxidative
stress, hypoxia, heat shock, nutrient deprivation, hyperosmolarity,
endoplasmic reticulum stress, and exposure to chemotherapeutic
drugs^[62]1,[63]2. In response to these challenges, a crucial adaptive
mechanism involves the formation of stress granules (SGs). SGs are
conserved cytoplasmic, non-membrane-bound ribonucleoprotein
compartments that dynamically assemble and disassemble through phase
separation and play a role in regulating mRNA storage, stability, and
translation during stress^[64]3,[65]4. Generally considered beneficial
for cell survival, SGs also contribute significantly to the
pathogenesis of various diseases, particularly cancer^[66]2. Recent
studies highlight that tumor cells exhibit a significantly higher
capacity to form SGs compared to normal cells^[67]5, emphasizing their
pronounced reliance on these organelles for survival, even during the
development of chemoresistance. Mitophagy, a specialized form of
autophagy, selectively removes damaged or dysfunctional mitochondria.
It plays a crucial role in maintaining mitochondrial quality control,
preventing oxidative damage, and ensuring cellular homeostasis during
stress^[68]6,[69]7. Although there has been a report suggesting that
SGs interact with mitochondria and down-regulate fatty acid β-oxidation
(FAO) during starvation stress^[70]8, research on the specific
relationship between SGs and mitophagy, as well as their combined
impact on cellular homeostasis and cancer progression, remains limited.
Emerging evidence suggests that SGs play a critical role in determining
cell fate by modulating mRNA stability under stress conditions^[71]9.
Several molecular mechanisms have been described for SGs in regulation
of mRNA stability. These include sequestration of RNA-binding proteins,
such as Hu-antigen R (HuR) and Zipcode-binding protein 1 (ZBP1), which
restrict their cytoplasmic functions and promote mRNA
degradation^[72]10, as well as the inhibition of nonsense-mediated
decay (NMD) through the concentration of essential NMD pathway
components like UPF1, SMG1, and UPF2^[73]11. Remarkably, only
approximately 10% of the total mRNA pool is recruited into stress
granules (SGs) and regulated during stress^[74]12. These selected mRNAs
exhibit distinct features, including extended lengths in both coding
and non-coding regions, specific sequence motifs such as
adenylate-uridylate (AU)-rich and guanine-cytosine (GC)-rich
elements^[75]13, and modifications like N6-methyladenosine
(m6A)^[76]14. Notably, mRNAs recruited into SGs, particularly those
with extended AU-rich motifs, predominantly belong to genes essential
for cell survival and proliferation^[77]13. Additionally, mRNAs bearing
m7G modifications specifically accumulate in SGs under stress,
impacting tumor cell resistance to chemotherapy^[78]9. However, the
precise molecular mechanisms by which SGs modulate mRNA stability under
stress conditions remain to be fully elucidated, and the selective
regulatory effects of SGs on mRNA stability are not yet fully
understood.
The core proteins that form SGs, along with their post-translational
modifications (PTMs), are critical for the assembly and functional
regulation of these structures^[79]15,[80]16. Accumulating evidence
highlights SUMOylation as a critical regulator of SG assembly and
disassembly. Studies report that SUMO molecules and SUMOylation enzymes
localize to SGs, and several SG-associated RNA-binding proteins (RBPs)
are identified as SUMOylation targets under stress
conditions^[81]17–[82]19. Knockdown of SAE1 (E1) or Ubc9 (E2)
significantly disrupts SG disassembly^[83]17. Furthermore, SUMOylation
facilitates SG disassembly by promoting ubiquitination via the
SUMO-targeted ubiquitin ligase (StUbL) pathway, linking these PTMs to
SG dynamics^[84]20. Poly(A)-Binding Protein Cytoplasmic 1 (PABPC1), an
mRNA poly(A) tail-binding protein, is considered a core marker protein
of SGs, along with G3BP1, TIA1, and others^[85]21. It typically
exhibits a diffuse cytoplasmic distribution under physiological
conditions^[86]22 while rapidly localizes to stress granules (SGs) in
response to cellular stress^[87]23. PABPC1 plays a crucial role in
regulating mRNA stability by binding to the mRNA poly(A) tail, which
protects the 3’ end of mRNA from deadenylation enzymes^[88]24,[89]25.
Conversely, PABPC1 can collaborate with deadenylases such as PAN2/3 and
the CCR4-NOT complex to promote mRNA degradation^[90]26,[91]27. Recent
studies demonstrate that PABPC1 undergoes ubiquitination at residues
K312, K512, K620, and K625 mediated by the E3 ubiquitin ligase MKRN3, a
modification shown to promote mRNA deadenylation and
degradation^[92]28. However, while PABPC1 is a core stress granule (SG)
component, it remains unclear whether additional PTMs, particularly
SUMOylation, regulate its functional dynamics under stress conditions.
In this study, we demonstrate that SUMOylation of PABPC1 significantly
enhances SG formation and promotes cancer cell survival by facilitating
mitophagy. This process involves recruiting and stabilizing U-rich
mRNAs within SGs under cellular stress. Specifically, various stress
stimuli robustly induce SUMOylation at the K512 site of PABPC1—a key
post-translational modification that strengthens its ability to
stabilize mRNAs involved in mitophagy. Consequently, this enhanced
stability leads to increase the expression of mitophagy-related genes,
ultimately elevating mitophagic activity, which is crucial for
maintaining cellular homeostasis and enhancing tumor cell resilience
under stress conditions. Mechanistically, SUMOylated PABPC1 interacts
with the SUMO-interacting motif (SIM) of the TIA1 protein. This
interaction selectively targets and recruits mRNAs with U-rich
sequences into stress granules, significantly improving their
stability. Notably, critical mitophagy-related genes, such as FUNDC1,
characterized by U-rich sequences in their 3’ UTRs, are effectively
recognized and protected by the PABPC1-SUMO-TIA1 complex during stress.
These findings underscore the critical role of PABPC1 SUMOylation as a
regulatory mechanism for mRNA stability and cellular adaptation to
stress, emphasizing its essential function in supporting tumor cell
survival in challenging environments.
Results
SUMOylation of PABPC1 is dynamically regulated by various stresses
To investigate whether PABPC1 undergoes SUMOylation in cells, we
transiently transfected Myc-PABPC1 and His-tagged SUMO1, SUMO2, or
SUMO3 into HEK-293T cells. We then pulled down His-SUMO-conjugated
PABPC1 using Ni^2+-NTA resin precipitation^[93]29. Western blotting
(WB) analysis revealed that PABPC1 was strongly modified by SUMO1 but
only weakly by SUMO2 or SUMO3 (Fig. [94]1a). Thus, in our follow-up
studies, we specifically investigated SUMO1 modification of PABPC1. To
further validate that PABPC1 is modified by SUMO1, we transfected
HA-PABPC1 alone or co-transfected it with His-tagged SUMO1, the
SUMO-conjugating enzyme E2 Flag-Ubc9, and the deSUMO enzyme SENP1
(Sentrin/SUMO-specific protease 1) into 293 T cells. Subsequently, we
pulled down His-SUMO1-conjugated PABPC1 using Ni^2+-NTA resin
precipitation and immunoblotted it with an anti-HA antibody. The
results showed that Ubc9 increased SUMOylated PABPC1, whereas
co-transfection with SENP1 greatly weakened this modification
(Fig. [95]1b). Secondly, we confirmed the SUMOylation of PABPC1 using
the method of denatured immunoprecipitation (IP)^[96]30. This method
allowed us to examine the crucial point that PABPC1 undergoes
endogenous modification by SUMO1. The results revealed that PABPC1 was
moderately modified by endogenous SUMO1 in 293 T cells upon SENP1
knockout using the CRISPR-Cas9 system (Fig. [97]1c, Supplementary
Fig. [98]1a). As expected, clear SUMOylation bands of PABPC1 were
detected in H1299 wild-type (WT) cells using both SUMO1 and PABPC1
antibodies. Notably, these bands were markedly diminished in PABPC1
knockout H1299 cells (Supplementary Figs. [99]1b-c), confirming the
specificity of the detected SUMOylation. Furthermore, to investigate
whether PABPC1 undergoes SUMOylation in vitro, we conducted a
prokaryotic SUMOylation assay in E. coli BL21 cells co-expressing
GST-PABPC1 along with the plasmid pT-E1E2S1^[100]31. This plasmid
simultaneously expresses two enzymes (E1 and E2) and SUMO1. Following
GST pull-down assay, immunoblotting using an anti-SUMO1 antibody
revealed that GST-PABPC1 co-transformed with pT-E1E2S1 was indeed
SUMOylated. The presence of SUMOylated bands was further confirmed by
detecting them with anti-GST and anti-PABPC1 antibodies (Fig. [101]1d).
These results conclusively demonstrate that PABPC1 undergoes
SUMOylation both in vivo and in vitro.
Fig. 1. PABPC1 is dynamically SUMOylated under stresses.
[102]Fig. 1
[103]Open in a new tab
a Myc-PABPC1 and Flag-Ubc9 were transfected with His-SUMO1, His-SUMO2
or His-SUMO3 into HEK-293T cells and the Ni^2+-NTA pulldown assay was
performed to detect SUMOylation of PABPC1. b HA-PABPC1, His-SUMO1 and
Flag-Ubc9 were transfected with or without EBG-SENP1 into HEK-293T
cells as indicated and SUMOylation of PABPC1 was accessed by Ni^2+-NTA
pulldown assay. c Denaturing immunoprecipitation (IP) was conducted to
evaluate endogenous SUMOylation of PABPC1 in HEK-293T^SENP1–/– Cells. d
GST pulldown assay was performed to validate the SUMOylation of PABPC1
in E. coli based in vitro system. e The levels of SUMOylation of PABPC1
were assessed after a time course treatment with AS in HEK-293T cells.
f Immunofluorescence staining was conducted to evaluate the formation
of PABPC1 (green) and G3BP1(red) foci at different time points after AS
treatment. Scale bar represented 10 μm, red arrows indicate foci formed
by PABPC1 and G3BP1. g The SUMOylation levels of PABPC1 were examined
in HEK-293T cells treated with AS for 1 h, followed by recovery for
various durations. h Immunofluorescence staining was performed to
monitor the changes in PABPC1 and G3BP1 foci following AS stimulation
for 1 h and subsequent recover for different time points in H1299
cells. Scale bar represented 10 μm, red arrows indicate foci formed by
PABPC1 and G3BP1. All results were shown with one representative image
from three independent experiments (a-h).
Next to investigate whether PABPC1 occurs SUMOylation in response to
stress conditions that induce stress granule formation, we transfected
293 T cells with His-SUMO1, Flag-Ubc9, and Myc-PABPC1. These cells were
then treated with 0.5 mM sodium arsenite (AS, NaAsO[2]), which induces
oxidative stress^[104]32,[105]33 and SG assembly^[106]34, for indicated
time before being harvested for a Ni^2+-NTA pull-down assay. The
results revealed that SUMOylation of PABPC1 increased upon AS
stimulation, reaching its peak within 1–3 h (h) (Fig. [107]1e).
Immunofluorescence (IF) staining demonstrated that PABPC1 formed the
most numerous and largest foci, which co-localized with SG marker
G3BP1, when its SUMO modification level was highest (Fig. [108]1f,
Supplementary Fig. [109]1d). Conversely, removal of AS led to reduced
SUMO modification of PABPC1, concomitantly with stress granule
disassembly (Fig. [110]1g), accompanied by the disappearance of PABPC1
foci (Fig. [111]1h, Supplementary Fig. [112]1e). These results suggest
that SUMOylation may regulate PABPC1 functions in response to cellular
stresses that induce SG formation.
Additionally, other stimuli, including heat stress, osmotic stress, and
glucose starvation, are known to induce stress granule formation. To
investigate PABPC1 SUMOylation in response to these stress conditions,
we conducted further experiments. Consistent with our findings from AS
treatment, PABPC1 SUMOylation increased upon exposure to heat shock
(Supplementary Figs. [113]1f, g), sorbitol treatment (Supplementary
Figs. [114]1h, i), and glucose starvation (Supplementary Figs. [115]1j,
k), while it decreased when these stresses were removed.
K512 is the major SUMOylation site of PABPC1
To identify the SUMOylation sites on PABPC1, we utilized the SUMOplot
tool ([116]http://www.abcepta.com/sumoplot) for predicting potential
SUMOylation sites (Supplementary Fig. [117]2a). Additionally, we
compared the potential SUMOylation sites on PABPC1 using mass
spectrometry data reported by Hendriks IA et al. (2017)^[118]35
(Supplementary Fig. [119]2b). The potential SUMO-site mutant plasmids,
alongside His-SUMO1 and Flag-Ubc9, were transfected into 293 T cells.
After 48 h, we assessed the SUMOylation of PABPC1 using a Ni^2+-NTA
pulldown assay. The results indicated that mutations at K167, K196,
K284, K324, K333, and K361 had no impact on PABPC1 SUMOylation
(Supplementary Figs. [120]2c–f, Fig. [121]2a). In contrast, the K512R
mutation led to the disappearance of a distinct band at approximately
120 kDa (Supplementary Fig. [122]2f), suggesting that this band likely
corresponds to SUMOylation at K512 of PABPC1. Consistently, targeted
mutagenesis of K512, K324, and K333 either individually or in
combination resulted in the complete absence of the band at ~120 kDa
(Fig. [123]2a). To validate these results, PABPC1-WT or PABPC1-K512R
was stably re-expressed in the PABPC1-knockdown (Fig. [124]2b) or
PABPC1-knockout (Fig. [125]2c) H1299 cell line, respectively. Following
SUMOylation analysis, PABPC1-K512R exhibited complete absence of the
band with a size of approximately 120 kDa. Denatured
immunoprecipitation (IP) followed by WB analysis showed that the
endogenous SUMOylation of PABPC1 was significantly reduced in the
PABPC1 K512R mutant (Fig. [126]2d). Additionally, pT-E1E2S1 was
co-transformed with either GST-tagged PABPC1-WT or PABPC1-K512R into E.
coli BL21 for a prokaryotic SUMOylation assay. Notably, the
GST-PABPC1-K512R mutant showed the complete disappearance of a major
specific SUMOylation-PABPC1 band (Fig. [127]2e). Furthermore, the
residue K512 of PABPC1 was confirmed as an endogenous SUMOylation site
in HeLa cells through the mass spectrometry (MS) analysis
(Supplementary Fig. [128]2g).
Fig. 2. K512 is the major SUMOylation site of PABPC1.
[129]Fig. 2
[130]Open in a new tab
a Identification of SUMOylation sites of PABPC1 in HEK-293T cells
transfected with plasmids containing individual or combined mutations
at K324, K333, and K512. Ni^2+-NTA pulldown assays detected SUMOylation
of PABPC1 in H1299 shPABPC1 (b) or H1299^PABPC1–/– (c) cells
re-expressing Myc-PABPC1-WT/K512R. d Denaturing immunoprecipitation
(IP) was conducted to evaluate endogenous SUMOylation of PABPC1 in
H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R. e GST pulldown
assay was performed to investigate SUMOylation of PABPC1-WT or
PABPC1-K512R in prokaryotic expression systems. f Myc-PABPC1-WT/K512R,
His-SUMO1, and Flag-Ubc9 were transfected into HEK-293T cells, and the
SUMOylation levels of PABPC1 were detected after treatment with AS for
1 and 3 h. g Protein structural modeling of PABPC1 and distribution of
its SUMOylation sites. * indicates the SUMO modification band at the
K512 site of PABPC1. All western blot data were shown with one
representative image from three independent experiments (a–f).
To determine whether the K512R mutation in PABPC1 exerts structural or
functional effects independent of its role in modulating SUMOylation,
we employed AlphaFold3 to predict the full-length structures of
PABPC1-WT and PABPC1-K512R, followed by structural alignment using
PyMOL. The K512R substitution resulted in minimal conformational
deviation (the root-mean-square deviation for the Cα atoms
(RMSD) = 0.555 Å) (Supplementary Fig. [131]2h), indicating negligible
impact on the overall protein architecture. Additional variants K512Q
(polar) and K512A (non-polar) showed similarly conserved structures
(Supplementary Fig. [132]2i), suggesting that alterations at K512 do
not affect PABPC1’s global folding. Further, EMSA results demonstrated
that both wild-type and K512R mutant PABPC1 efficiently bound to A90
RNA, forming stable complexes, and importantly, the K512R mutation did
not impair the protein’s ability to bind to mRNA poly(A) tails
(Supplementary Fig. [133]2j). Together, these results demonstrate that
the K512R mutation specifically alters PABPC1 SUMOylation without
affecting its structural integrity or RNA-binding function.
To further elucidate SUMOylation at K512 of PABPC1 in response to
stress, both PABPC1-WT and PABPC1-K512R were exposed to AS treatment
for 1 and 3 h. The results of Ni^2+-NTA pull down assay revealed a
markedly increase in SUMOyaltion of PABPC1-WT following AS treatment,
whereas SUMOylation of PABPC1-K512R was not induced under the same
conditions (Fig. [134]2f). Similarly, under conditions such as heat
shock, sorbitol treatment, and glucose deprivation, SUMOylation of
PABPC1-WT was markedly elevated, while the 120-kDa SUMOylated band was
absent in PABPC1-K512R (Supplementary Figs. [135]2k-m). These finding
conclusively established K512 as a crucial SUMOylation site of PABPC1,
particularly under stress.
Previous studies have indicated that K512 serves as one of the
ubiquitination sites on PABPC1, influencing its binding to poly(A) and
consequently affecting mRNA stability^[136]28. In our ubiquitination
assay using 293 T cells, we co-transfected Flag-Ub along with either
Myc-PABPC1-WT or Myc-PABPC1-K512R. The results demonstrated that
PABPC1-WT underwent ubiquitination, while the K512R mutation weakened
its ubiquitination to some extent (Supplementary Fig. [137]2n),
confirming K512 as one of the ubiquitination sites on PABPC1. However,
to investigate whether ubiquitination at K512 responds to stress, we
conducted the same experiments with PABPC1-WT and PABPC1-K512R,
treating them with AS. Interestingly, we observed a significant
upregulation in the ubiquitination of PABPC1-WT during and after
exposure to AS. Surprisingly, this upregulation was not hindered by the
K512R mutation (Supplementary Fig. [138]2o), suggesting that K512 of
PABPC1 primarily functions as a SUMO modification site in response to
stress. Additionally, the K512 site is located within the linker region
of PABPC1 (Fig. [139]2g), which is known for its high degree of
disorder often associated with stress-triggered phase separation
functions^[140]36. This structural observation suggests that
SUMOylation of PABPC1 at the K512 site could impact the regulation of
SGs, providing a perspective on PABPC1 function under stress
conditions.
PABPC1 SUMOylation promotes SG formation
Previous studies demonstrated that SUMO modification plays a role in
stress granule (SG) disassembly^[141]20,[142]37. To further investigate
the impact of PABPC1 SUMOylation on SG formation during stress, we
utilized CRISPR/Cas9 technology to create a SENP1 knockout (SENP1^–/–)
in the H1299 cell line (Supplementary Fig. [143]3a). Immunofluorescence
(IF) analysis revealed a widespread cytoplasmic distribution of PABPC1
and G3BP1 proteins under normal condition. However, exposure to
oxidative stress (induced by arsenite treatment) led to the rapid
coalescence of these proteins into fluorescent foci, indicative of SG
formation. Interestingly, SENP1-deficient cells exhibited an enhanced
capacity for SG formation and more efficient disassembly upon recovery
from stress conditions (Fig. [144]3a). These findings highlight the
crucial role of SUMOylation in both SG assembly and subsequent
disassembly.
Fig. 3. PABPC1 SUMOylation promotes stress granule formation.
[145]Fig. 3
[146]Open in a new tab
a Immunofluorescence staining of PABPC1 and G3BP1 in H1299^SENP1+/+ or
H1299^SENP1–/– cells under AS-mediated stress condition. Scale bar
represented 20 μm and statistical graphs showing the number of SGs. b
Immunofluorescence staining of GFP-tagged PABPC1 and G3BP1 in
H1299^PABPC1–/– cells re-expressing GFP-PABPC1-WT/K512R treated with AS
for difference concentration. SGs foci were quantified by CellProfiler
software and presented with dot graph. Scale bar represented 20 μm. c
Immunofluorescence staining of GFP-tagged PABPC1 and G3BP1 in
H1299^PABPC1–/– cells re-expressing GFP-PABPC1-WT/K512R treated with AS
for 1 h and recovery for indicated time. Statistical graphs showing the
number of SGs. Scale bar represented 20 μm. d Immunofluorescence
staining of GFP-tagged PABPC1 and G3BP1 in H1299^PABPC1-/- cells
re-expressing GFP-PABPC1-WT/K512R treated with AS for 1 h and 2-D08
(pan SUMOylation inhibitor) for difference concentration for 24 h.
Statistical graphs showing the number of SGs. Scale bar represented
20 μm. e FRAP analysis of GFP-tagged PABPC1-WT/K512R in H1299^SENP-/-
cells. The dashed circle with 0.8 μm diameter inside a large droplet
was selected for photobleaching. Relative fluorescence intensity
plotted as line graph over time. Mobile fraction was showed as bar
graph, mean ± SD, n = 3 independent droplets. f Workflow for the
isolation of PABPC1 interacting SG proteins. Volcano plots showing the
differences in PABPC1-bound SG proteins in H1299^PABPC1–/– cells
re-expressing Myc-PABPC1-WT/K512R after 1 h of AS treatment (g) and a
90 min recovery (h). i Gene Ontology (GO) pathway enrichment analysis
was performed using proteins that are 1.5-fold up-regulated in binding
with PABPC1-WT under AS treatment, P-values were calculated by DAVID
bioinformatics tools using Fisher’s Exact Test. The data in (a–e) were
shown as one representative image from three independent experiments.
For Immunofluorescence staining, Statistical data are presented as
mean ± SD, n ≥ 100 cells; P-values were calculated using two-tailed
Student’s t test (a–d). For mass spectrometry analysis, two samples
each group, P-values were calculated by two-tailed Student’s t test (g,
h).
To explore the specific regulatory role of SUMOylation at K512 of
PABPC1 in SG dynamics, we first knocked down PABPC1 using shRNA in HeLa
cells and performed immunofluorescence (IF) staining following AS
treatment for 1 h to assess the role of PABPC1 in SG formation. The
staining results showed that PABPC1 depletion did not impair SG
formation (Supplementary Fig. [147]3b), consistent with prior findings
that the loss of individual SG proteins does not always lead to
complete suppression of SG assembly^[148]38. Second, we generated
H1299^PABPC1–/– and HeLa^PABPC1–/– cell lines re-expressing either
GFP-PABPC1-WT or GFP-PABPC1-K512R (Supplementary Figs. [149]3c–e) and
conducted IF staining following AS treatment for 1 h. The results
revealed that the K512R mutation in PABPC1 attenuated SG formation
compared to PABPC1-WT (Fig. [150]3b, Supplementary Fig. [151]3f).
Interestingly, during recovery from AS exposure, the difference in SG
disassembly between PABPC1-WT and PABPC1-K512R expressing cells was
minimal (Fig. [152]3c, Supplementary Fig. [153]3g). These findings
suggest that while SUMOylation at K512 of PABPC1 significantly promotes
SG assembly, it does not significantly affect SG disassembly.
To further confirm the role of K512 SUMOylated PABPC1 in SG formation,
we overexpressed GFP-tagged PABPC1-WT or PABPC1-K512R in H1299 cells
and H1299 SENP1 knockout (H1299^SENP1–/–) cells. After AS treatment for
1 h and subsequent IF analysis, we found that PABPC1-WT overexpression
resulted in a significantly greater number of SGs compared to cells
expressing PABPC1-K512R, in both H1299 and H1299^SENP1–/– cells.
Notably, SG formation was more pronounced in H1299^SENP1–/– cells
compared to H1299 cells (Supplementary Fig. [154]3h). Furthermore, we
treated H1299^PABPC1–/– cells re-expressing either PABPC1-WT or
PABPC1-K512R with 2-D08, a pan-SUMOylation inhibitor, to globally
suppress SUMOylation. IF analysis showed a marked decrease in SG
formation in PABPC1-K512R cells compared with PABPC1-WT cells under AS
treatment. Treatment with 2-D08 further impaired SG formation in both
PABPC1-WT and PABPC1-K512R cells. Notably, at a 150 µM concentration of
2-D08, the difference in SG formation between PABPC1-WT and
PABPC1-K512R cells was no longer significant, likely due to the
complete suppression of PABPC1 SUMOylation, which abolished its
regulatory role in SG assembly (Fig. [155]3d). These results highlight
the critical regulatory role of SUMOylation in SG dynamics, with PABPC1
SUMOylation acting as a positive modulator of this process. Moreover,
fluorescence recovery after photobleaching (FRAP) assays revealed that
PABPC1-WT displayed faster fluorescence recovery compared to the K512R
mutant (Fig. [156]3e), suggesting that SUMOylation at K512 enhances the
dynamic exchange of PABPC1 within phase-separated condensates, thereby
promoting SG formation.
Next, we isolated SGs in H1299^PABPC1–/–cells re-expressing
Myc-PABPC1-WT or Myc-PABPC1-K512R for mass spectrometry analysis on
interacting proteins with PABPCl in SGs (Fig. [157]3f). The effective
isolation of SGs was confirmed by detecting key SG proteins, including
G3BP1 and TIA1, through Co-immunoprecipitation (Co-IP) using an
anti-Myc antibody specific for PABPC1 (Supplementary Fig. [158]3i).
Mass spectrometry revealed a significant decrease in the number of
interacting proteins with the PABPC1-K512R mutant compared to PABPC1-WT
under arsenite treatment for 1 h (Fig. [159]3g, Supplementary
data [160]2). Interestingly, after AS treatment for 1 h followed by
removal and a 90-minute recovery period, there were no significant
differences in the number of interacting proteins between PABPC1-WT and
PABPC1-K512R in SGs (Fig. [161]3h, Supplementary data [162]2). Further
analysis showed that during recovery, the reduction in SG proteins
associated with either PABPC1-WT (Supplementary Fig. [163]3j,
Supplementary data [164]2) or PABPC1-K512R (Supplementary Fig. [165]3k,
Supplementary data [166]2) was similar, suggesting that SUMOylation at
K512 of PABPC1 does not impact SG disassembly. Based on previous
immunofluorescence data, we conclude that PABPC1 SUMOylation enhances
its aggregation with other SG proteins, promoting SG formation under
stress conditions without affecting subsequent disassembly.
Additionally, we conducted a Gene Ontology (GO) pathway enrichment
analysis on proteins that exhibit 1.5-fold upregulation in their
binding affinity with PABPC1-WT compared to PABPC1-K512R during AS
treatment. The analysis revealed a significant enrichment of these
proteins in pathways related to RNA stability regulation
(Fig. [167]3i). This finding suggests that SUMOylation at K512 of
PABPC1 may play a role in modulating RNA stability under stress.
SUMOylation of PABPC1 promotes mRNA stability under stress
Given that PABPC1 is SUMOylated at K512 to enhance SG formation, we
investigated whether this SUMOylation also regulates mRNA stability. To
address this, we performed 4-Thiouridine (4sU) Pulse-Chase sequencing
(4sU-Seq) in H1299^PABPC1-/-cells re-expressing Myc-PABPC1-WT or
Myc-PABPC1-K512R to examine mRNA stability genome-wide. After labeling
the cells with 4sU, we removed the label and treated them with either
PBS or 0.5 mM AS. Samples were collected at 0, 3, and 6 h
post-treatment for biotin labeling and RNA isolation, followed by RNA
sequencing (Fig. [168]4a, Supplementary data [169]3). Half-life
determination revealed a mild effect on mRNA half-lives between
PABPC1-WT and PABPC1-K512R groups under normal conditions, with the
K512R mutation showing a slight increase in mRNA half-life
(Fig. [170]4b). The distribution of these half-lives across both groups
did not exhibit marked differences (Fig. [171]4c), and their median
half-life values were comparable (Supplementary Fig. [172]4a). However,
under stress conditions, we observed a pronounced reduction in mRNA
half-life and a shorter half-life distribution in the PABPC1-K512R
cells compared to the PABPC1-WT group (Fig. [173]4d, e), with a notably
lower median half-life value (Supplementary Fig. [174]4b).
Fig. 4. SUMOylation of PABPC1 promotes mRNA stability under stress.
[175]Fig. 4
[176]Open in a new tab
a Flowchart of the 4sU pulse-chase sequencing (4sU-Seq) process.
Cumulative distribution plot (b) and frequency distribution plot (c) of
mRNA half-life in H1299^PABPC1–/– cells re-expressing
Myc-PABPC1-WT/K512R under normal condition. Cumulative distribution
plot (d) and frequency distribution plot (e) of mRNA half-life in
H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R under
AS-mediated stress condition. f KEGG pathway analysis of genes in the
4sU-Seq data that showing a twofold increase in half-life in
H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT under stress
condition, P-values were calculated by DAVID bioinformatics tools using
Fisher’s Exact Test. g List of enriched genes in the Mitophagy pathway.
h, i 4sU-RT-qPCR was performed to detect the stability of FUNDC1,
BECN1, and BNIP3L mRNA in H1299^PABPC1–/– cells re-expressing
Myc-PABPC1-WT/K512R under normal and stress conditions. Decay graphs
were generated by applying the one-phase decay model. For Cumulative
fraction analysis, P-values were calculated using a two-sided
Mann–Whitney U test (b, d). For 4sU-RT-qPCR, data were presented as
mean ± SD, n = 3 biologically independent replicates, P-values were
determined by one-side sum-of-squares F test (h–j).
Subsequently, we stratified the sequencing data to assess differences
in mRNA half-lives between PABPC1-WT and PABPC1-K512R cells under both
normal and stress conditions. Our analysis revealed that AS treatment
shortened mRNA half-lives compared to normal conditions. Remarkably,
this reduction in mRNA half-life was more pronounced in the
PABPC1-K512R groups following AS exposure (Supplementary Fig. [177]4c,
d). This trend implies a superior mRNA stabilization capability in
PABPC1-WT cells when subjected to stress (Supplementary Fig. [178]4e).
These observations indicate that SUMOylation of PABPC1 under stress
conditions contributes to enhanced mRNA stability. Interestingly, under
normal conditions, both the wild-type and the SUMO1-site mutant of
PABPC1 demonstrate comparably low levels of SUMOylation, resulting in
negligible differences in their impact on mRNA stability.
To further investigate the biological implications of PABPC1
SUMOylation under stress conditions, we conducted KEGG pathway
enrichment analysis and Gene Ontology (GO) analysis on transcripts with
a twofold decrease in mRNA half-life in the K512R mutant group. The
KEGG pathway analysis revealed that these transcripts were
significantly enriched in pathways such as Pathways in Cancer, Wnt
signaling, and Mitophagy (Fig. [179]4f). Notably, both KEGG and GO
analyses consistently highlighted that the transcripts stabilized by
PABPC1 SUMOylation were predominantly associated with the mitophagy
pathway (Supplementary Fig. [180]4f). Furthermore, 11 genes with
altered mRNA half-lives were found to be associated with the mitophagy
pathway, including key mitophagy-related genes such as FUNDC1, BNIP3L,
and BECN1 (Fig. [181]4g).
Previous studies have demonstrated that stress conditions, such as
nutrient deprivation and hypoxia, can activate mitophagy in tumors to
eliminate damaged mitochondria and recycle metabolic intermediates,
thereby promoting tumor cell survival^[182]39,[183]40. The upregulation
of mitophagy-related proteins, including FUNDC1, NIX, and ARIH1, has
been shown to protect tumor cells from chemotherapy-induced cell
death^[184]41–[185]43. These findings suggest that PABPC1 SUMOylation
may regulate the stability of mitophagy-related transcripts, thereby
influencing mitophagy and contributing to the adaptive responses of
tumor cells under stress conditions. To validate these observations, we
evaluated the mRNA stability of genes enriched in mitophagy pathway in
PABPC1-WT cells and PABPC1-K512R cells using 4sU-RT-qPCR analysis. As
expected, the mRNA half-life of FUNDC1, BNIP3L, and BECN1 were markedly
decreased in the PABPC1-K512R group compared to the PABPC1-WT under
stress conditions. Interestingly, under normal conditions, the mRNA
half-life of these three genes showed little difference between
PABPC1-WT and PABPC1-K512R groups (Fig. [186]4h–j). Notably, these
three transcripts were partially localized in SG core fraction
(Supplementary Fig. [187]4g), suggesting that their stability may be
protected by SUMOylated PABPC1 within SGs under stress conditions.
Additionally, the mRNA half-lives of other mitophagy-related genes,
such as MITF, FOXO3, and HRAS, were also reduced in the PABPC1-K512R
group under stress conditions (Supplementary Fig. [188]4h–j).
Collectively, these results support the proposition that elevated
SUMOylation of PABPC1 under stress conditions plays a critical role in
regulating mRNA stability.
SUMOylation of PABPC1 enhances its affinity with mRNA under stress
Previous research has shown that mRNA stability is largely influenced
by the binding affinity of PABPC1 to the poly(A) tail^[189]28.
Therefore, we propose that under stress conditions, the molecular
mechanism by which SUMOylated PABPC1 regulates mRNA stability is
intricately linked to its RNA-binding affinity. To prove this, we
employed RNA Immunoprecipitation sequencing (RIP-Seq) using an anti-Myc
antibody in H1299^PABPC1-/- cells re-expressing either Myc-PABPC1-WT or
Myc-PABPC1-K512R. Firstly, we evaluated the binding capability of
PABPC1 to mRNA under both normal condition and oxidative stress induced
by AS treatment. The RIP-Seq results revealed an enhanced binding of
PABPC1 to a larger number of transcripts under stress (P = 7.0E − 32,
Mann–Whitney U test) (Fig. [190]5a, Supplementary data [191]4). We
further investigated whether PABPC1-WT and PABPC1-K512R exhibited
differences in mRNA affinity under stress conditions. Cumulative
distribution analysis demonstrated that the K512R mutation
significantly reduced the binding of PABPC1 to mRNA (P = 3.0E − 31,
Mann–Whitney U test) (Fig. [192]5b, Supplementary data [193]4). By
analyzing RIP-Seq data from WT-NT (PABPC1-WT group under normal
condition), WT-AS (PABPC1-WT group under stress condition), and
K512R-AS (PABPC1-K512R group under stress condition), we identified
5623 transcripts (Fig. [194]5c). Notably, these mRNA transcripts
predominantly exhibited decreased binding in the K512R-AS group, while
also showing reduced binding in the WT-NT group compared to the WT-AS
group (Fig. [195]5d). To corroborate the aforementioned conclusion that
SUMOylation of PABPC1 enhances its mRNA affinity based on RIP-Seq data,
we conducted oligo(dT) pull-down experiments. In 293 T cells, we
co-transfected His-SUMO1, Flag-Ubc9, Myc-PABPC1-WT or K512R, and SENP1.
Subsequently, we pulled down PABPC1 bound to poly(A) RNA using
oligo(dT) beads. Our results clearly indicated that SUMOylation clearly
enhances the binding of PABPC1 to mRNA. In contrast, the K512R mutation
notably weakens this interaction, an effect further intensified by
SENP1 (Fig. [196]5e).
Fig. 5. SUMOylation of PABPC1 at K512 enhances its affinity with mRNA under
stress.
[197]Fig. 5
[198]Open in a new tab
a Cumulative fraction analysis of RIP-Seq for mRNA transcripts bound to
PABPC1 in H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT under
normal and AS-mediated stress conditions. Boxes indicate the median,
25th and 75th percentiles, the whiskers denote the minima and maxima. b
The cumulative distribution diagram shows the mRNA transcripts bound to
PABPC1 in H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R
treated with AS. Boxes indicate the median, 25th and 75th percentiles,
the whiskers denote the minima and maxima. c Venn diagram analysis of
mRNA transcripts bound to PABPC1 in the three groups of cells used in
(a) and (b). WT_NT: transcripts bound to PABPC1 in H1299^PABPC1–/–
cells re-expressing Myc-PABPC1-WT under normal condition; WT_AS:
transcripts bound to PABPC1 in H1299^PABPC1-/- cells re-expressing
Myc-PABPC1-WT under AS-mediated stress condition; K512R_AS: transcripts
bound to PABPC1 in H1299^PABPC1–/– cells re-expressing Myc-PABPC1-K512R
treated with AS. d Scatter plots analysis of mRNA transcripts with at
least 1.5-fold alteration in binding to PABPC1 in the intersection of
the data from (c). e oligo(dT) pulldown was performed in HEK-293T cells
transfected with Myc-PABPC1-WT/K512R, His-SUMO1, Flag-Ubc9 and
EBG-SENP1 to detect the effect of SUMOylation on the binding of PABPC1
to poly(A) RNAs. f RIP-qPCR detecting the binding of FUNDC1, BNIP3L,
and BECN1 mRNAs to PABPC1 in H1299^PABPC1-/- cells re-expressing
Myc-PABPC1-WT/K512R under normal and AS-mediated stress conditions.
Cumulative distribution plot (g) and Heatmap (h) displaying mRNA
half-life for transcripts recruited to PABPC1 with more than 1.5-fold
up-regulated in WT_AS group compared to K512R_AS group; Boxes indicate
median, 25th and 75th percentiles, and whiskers extend to 1.5 times the
interquartile range (g). For Cumulative fraction analysis, P-values
were calculated using a two-sided Mann–Whitney U test (a, b, g). For
RIP-qPCR, presented as mean ± SD, n = 3 biologically independent
replicates, P-values were determined by one-way ANOVA (f). Western blot
data were shown with one representative image from three independent
experiments (e).
The KEGG pathway enrichment analysis of genes with a 1.5-fold increase
in binding to PABPC1-WT in RIP-Seq revealed significant enrichment in
the mitophagy pathway (Supplementary Fig. [199]5a), consistent with our
previous 4sU-Seq pathway enrichment analysis. Furthermore, a combined
analysis of RIP-Seq and 4sU-seq data identified 367 transcripts with
enhanced binding to PABPC1 and increased mRNA half-life in PABPC1-WT
cells (Supplementary Fig. [200]5b). The KEGG pathway enrichment
analysis of these genes also demonstrated effective enrichment in the
mitophagy pathway (Supplementary Fig. [201]5c). Subsequently, we
employed RIP-RT-qPCR in H1299 and HeLa stable cell lines to examine the
binding between PABPC1 and mitophagy-related genes FUNDC1, BNIP3L, and
BECN1. The results showed no significant difference in the binding of
these three genes between PABPC1-WT and K512R under normal conditions.
However, under stress conditions, PABPC1-WT exhibited significantly
enhanced binding to these genes, while the binding in K512R was
substantially lower compared to the WT group (Fig. [202]5f,
Supplementary Fig. [203]5d). In conclusion, our research highlights
that SUMOylation at K512 of PABPC1 significantly enhances its
interaction with mRNAs, such as FUNDC1, BNIP3L, and BECN1, under stress
conditions.
Furthermore, we investigated whether SUMOylation enhances PABPC1’s
affinity for mRNA, thereby regulating mRNA stability. The RNA stability
analysis revealed a 1.5-fold increase in binding to the WT-AS group
under stress conditions, resulting in longer half-life (Fig. [204]5g,
h). This suggests that SUMOylation of PABPC1 influences mRNA stability
by affecting its binding to mRNAs. Collectively, these findings
underscore the critical role of PABPC1 SUMOylation in enhancing mRNA
binding and contributing to mRNA stabilization.
The PABPC1-SUMO-TIA1 complex recruits U-rich mRNAs to SGs
To explore whether mRNAs regulated by SUMOylated PABPC1 under stress
conditions display specific characteristic features, we analyzed
motif-based sequences using the MEME Suite tools^[205]44. Our
investigation revealed that transcripts exhibiting enhanced affinity
for PABPC1-WT in the RIP-Seq data were predominantly characterized by
U-Rich and A-Rich (poly(A)) motifs (Supplementary data [206]5). The
presence of poly(A) characteristics likely reflects PABPC1’s inherent
binding affinity toward mRNA’s poly(A) tail. Analysis of the 3ʹ
untranslated regions (3ʹ UTRs) of mRNAs, whose stability is influenced
by SUMOylation of PABPC1, revealed a prevalence of U-Rich motifs in
these regions. Remarkably, transcripts concurrently regulated by
SUMOylated PABPC1 in both the RIP-Seq and 4sU-Seq datasets also
exhibited these U-Rich features (Fig. [207]6a). Importantly, the
cumulative distribution plot revealed that the K512R mutation
significantly suppresses the interaction between PABPC1 and U-rich
mRNAs under stress conditions (Fig. [208]6b). Furthermore, analysis of
the proportion of U-rich mRNAs revealed that 8.93% of the transcripts
displayed reduced binding to SUMOylated PABPC1, whereas 13.86% of the
transcripts showed increased binding to SUMOylated PABPC1
(Supplementary Fig. [209]6a), implying that SUMOylation at K512
enhances the interaction of PABPC1 with U-rich mRNAs under stress
conditions. Taken together, these findings suggest that transcripts
modulated by SUMOylation of PABPC1 under stress conditions are
predominantly characterized by U-rich sequences, shedding light on the
precise regulatory roles of PABPC1 in stress response.
Fig. 6. The PABPC1-SUMO-TIA1 complex recruits U-rich mRNAs to SGs.
[210]Fig. 6
[211]Open in a new tab
a Motif discovery of transcripts in sequencing data using MEME
software. Top: Motif analysis of transcripts exhibiting a 1.5-fold
increase in binding to PABPC1-WT compared to PABPC1-K512R in RIP-Seq
data under stress condition; Middle: Motif analysis of transcripts with
a twofold up-regulated in mRNA half-life in H1299^PABPC1–/– cells
re-expressing Myc-PABPC1-WT under stress condition, based on 4sU-Seq
data; Bottom: Motif analysis of transcripts identified at the
intersection of the above two datasets. b Cumulative fraction analysis
of U-rich mRNA transcripts bound to PABPC1 in H1299^PABPC1–/– cells
re-expressing Myc-PABPC1-WT under AS-mediated stress conditions, based
on RIP-Seq data. c Co-Immunoprecipitation (Co-IP) detecting the
interaction between PABPC1 and TIA1 in HEK-293T cells co-trasfected
with His-SUMO1, Flag-Ubc9, and EBG-SENP1. d Co-Immunoprecipitation
(Co-IP) accessing the interaction between PABPC1 and TIA1 in
H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R under
AS-mediated stress condition. e Co-Immunoprecipitation (Co-IP) showing
the potential interaction between PABPC1 and several SIM mutated TIA1
in HEK-293T cells co-transfected with His-SUMO1 and Flag-Ubc9 plasmids.
f RIP-qPCR detecting the binding of FUNDC1, BNIP3L, and BECN1 mRNAs to
TIA1 under normal and stress conditions. g In vitro poly(U) RNA binding
assay to detect the directly interaction of purified His-GFP-PABPC1 or
His-GFP-PABPC1 + pE1E2S1, GST-TIA1, and Bio-poly(U) RNA. h Schematic
diagram of TIA1 and PABPC1 binding to FUNDC1 mRNA. For Cumulative
fraction analysis, P-values were calculated using a two-sided
Mann–Whitney U test; in box plots, the lines represent the median, 25th
and 75th percentiles, the whiskers denote the minima and maxima (b).
For RIP-qPCR, data were presented as mean ± SD, n = 3 biologically
independent replicates, P-values were determined by two-tailed unpaired
t test (f). Western blot data were shown with one representative image
from three independent experiments (c–e, g).
Given that PABPC1 is a classic mRNA poly(A) tail-binding protein with
generally non-specific interactions, we speculated that under stress
conditions, the specific regulation of U-rich mRNA by SUMOylated PABPC1
might involve a U-rich sequence-binding protein. Interestingly, recent
studies have demonstrated that under stress conditions, the RNA-binding
protein TIA1 preferentially interacts with alternative 3ʹ UTR sequences
enriched in U-rich motifs, a process linked to stress granule formation
and mRNA decay^[212]45. Beyond its well-known function in binding
AU-rich motifs within mRNA 3ʹ UTRs to regulate translation^[213]46,
TIA1 has also been implicated in the recruitment of RNA to SGs, which
are crucial for the cellular stress response^[214]23. Building on these
findings, we proposed that under stress conditions, SUMOylated PABPC1
might interact with TIA1, facilitating the preferential binding to
U-rich mRNAs and modulating their stability.
To confirm this, we conducted co-transfection experiments in 293 T
cells using a combination of HA-TIA1, Myc-PABPC1-WT or K512R,
His-SUMO1, Flag-Ubc9, and EBG-SENP1. The co-immunoprecipitation
followed by Western blot (CO-IP/WB) results clearly demonstrated that
SUMOylation enhanced the interaction between PABPC1 and TIA1.
Importantly, this interaction was impaired by the K512R mutation and
further reduced upon SENP1 expression (Fig. [215]6c). Consistently, in
H1299 stable cell lines, elevated SUMO modification levels were
associated with increased PABPC1-TIA1 interaction (Supplementary
Fig. [216]6b). Further analysis following AS treatment of H1299 stable
cell lines revealed a more pronounced interaction between endogenous
TIA1 and PABPC1-WT compared to the K512R variant (Fig. [217]6d).
Immunofluorescence (IF) studies corroborated this observation,
demonstrating the co-localization of TIA1 with PABPC1 within SGs under
stress (Supplementary Fig. [218]6c). Collectively, these findings
suggest that stress-induced SUMOylation of PABPC1 facilitates its
interaction with TIA1.
To further elucidate the interaction mechanism between TIA1 and PABPC1,
we utilized the JASSA online prediction tool
([219]http://www.jassa.fr/) to identify SUMO-interacting motifs (SIMs)
in TIA1 (Supplementary Fig. [220]6d). Subsequent alanine mutations in
these domains, followed by CO-IP/WB assays, revealed that the
RRM1-domain SIM1/SIM2 mutation partially reduced TIA1’s binding to
SUMOylated PABPC1, while the RRM3-domain SIM3 mutation nearly abolished
the interaction (Fig. [221]6e). This finding indicates that TIA1
interacts with SUMOylated PABPC1 predominantly through its SIM3.
Furthermore, we employed Electrophoretic Mobility Shift Assay (EMSA) to
validate the interaction between TIA1 and U-rich RNA. Our results
demonstrated that TIA1 directly binds to U-rich RNAs in vitro, with
binding strength increasing in a concentration-dependent manner
(Supplementary Fig. [222]6e). Notably, Structural alignment revealed
that the SIM1&2 mutations induced minimal perturbations in the overall
conformation of the TIA1 RRM1 motif (RMSD = 1.253 Å). Nevertheless,
subtle local conformational shifts were detected, particularly within
α-helical regions and adjacent flexible loops, indicating SIM mutations
within the RRM1 motif of TIA1 may have functionally relevant effects
(Supplementary Fig. [223]6f). In contrast, the SIM3 mutation, located
within the RRM3 motif of TIA1, had a negligible effect on the global
structure of the TIA1 RRM3 domain (RMSD = 0.169 Å) (Supplementary
Fig. [224]6g). Consistent with these structural observations, mutations
in SIM1 & 2 impaired the binding of TIA1 to poly (U) RNA, while the
mutation in SIM3 had little effect on the interaction between TIA1 and
poly (U) RNA (Supplementary Fig. [225]6h). Sequence analysis revealed
that the mRNA 3’ UTRs of mitophagy-related genes FUNDC1, BNIP3L, and
BECN1 contain U-rich sequences (Supplementary Fig. [226]6i). RIP-qPCR
experiments indicated that AS treatment significantly enhances the
binding of TIA1 to these three transcripts (Fig. [227]6f). In parallel,
we investigated the effects of TIA1 expression on the stability and
expression of U-rich RNAs. qPCR results revealed that knocking down
TIA1 in H1299 cells did not significantly alter the steady-state levels
or stability of FUNDC1, BNIP3L, and BECN1 compared to the control
group, suggesting that TIA1 expression levels have little to no effect
on the stability and expression of U-rich RNAs (Supplementary
Fig. [228]6j–n). The above results suggest that SUMOylation facilitates
the interaction between the covalently attached SUMO1 moieties on
PABPC1 and the SIMs of TIA1, thereby promoting the binding to U-rich
mRNA under stress condition. To further validate this hypothesis, we
conducted in vitro poly(U) RNA binding assays to assess the direct
interaction between SUMOylated PABPC1, TIA1, and U-rich RNA.
Recombinant His-GFP-PABPC1, SUMO1-modified PABPC1 (His-GFP-PABPC1 and
pT-E1E2S1 cotransfected), and GST-tagged TIA1 proteins were purified
and incubated with poly(U) RNA. The protein complexes were captured
using Ni²⁺-NTA resin precipitation and analyzed by Western blotting and
Northern blotting (Fig. [229]6g). The results confirmed that SUMO
modification enhances the binding of PABPC1 to both TIA1 and poly(U)
RNA. Taken together, these findings support a model in which
stress-induced SUMOylation facilitates the formation of the
PABPC1-SUMO1-TIA1 complex, which specifically recognizes and stabilizes
U-rich mRNAs, such as FUNDC1, by recruiting them to SGs and protecting
them from degradation (Fig. [230]6h).
SUMOylation of PABPC1 promotes cell survival by enhancing mitophagy under
stress
Above findings suggested that SUMOylation of PABPC1 specifically
enhances the stability of U-rich mRNAs, including critical genes
involved in mitophagy—a highly conserved cellular process that
maintains cellular homeostasis by eliminating dysfunctional or excess
mitochondria through autophagy^[231]6. Numerous studies have
demonstrated the significant role of mitophagy in tumorigenesis and
cancer cell survival within the tumor microenvironment^[232]47,[233]48.
Additionally, accumulating evidence highlights the role of SGs in
promoting cancer cell survival^[234]49. Therefore, we propose that
SUMOylation of PABPC1 acts as a bridge between SGs and mitophagy,
promoting their synergistic regulation of cellular homeostasis and
influencing cancer progression under stress. To address this, we
investigated whether SUMOylation of PABPC1 is linked to cancer cell
survival under stress. Cell proliferation assays revealed no
significant difference in cancer cell growth between PABPC1-WT and
PABPC1-K512R under normal conditions (Fig. [235]7a). However, when
H1299 and HeLa stable cell lines were exposed to AS-induced stress,
cell viability assays showed a pronounced reduction in cell vitality in
the PABPC1-K512R cells compared to PABPC1-WT cells (Fig. [236]7b,
Supplementary Fig. [237]7a). Clonogenic survival assays further
supported this observation, showing increased colony formation in
PABPC1-WT cells and fewer colonies in the K512R cells under AS
treatment (Fig. [238]7c, Supplementary Fig. [239]7b).
Fig. 7. SUMOylation of PABPC1 enhance mitophagy to promotes cell survival
under stress condition.
[240]Fig. 7
[241]Open in a new tab
a CCK-8 assay assessing proliferation of H1299^PABPC1–/– cells
re-expressing Myc-PABPC1-WT/K512R under normal condition. CCK-8 (b) and
plate colony formation (c) assay evaluating cell viability and
clonogenic ability of H1299^PABPC1–/– cells re-expressing
Myc-PABPC1-WT/K512R upon AS treatment. d Cell viability of H1299 and
H1299^SENP1–/– cells overexpressing Myc-PABPC1-WT/K512R cells treated
with 20 μM AS for 48 h. e Cell viability of H1299^PABPC1–/– cells
re-expressing Myc-PABPC1-WT/K512R treated with 20 μM AS and different
concentration of 2-D08 for 48 h. f qPCR quantification of FUNDC1,
BNIP3L, and BECN1 mRNA levels in H1299^PABPC1–/– cells re-expressing
Myc-PABPC1-WT/K512R under AS-induced stress. g Western blotting
analysis of FUNDC1, BNIP3L, and BECN1 protein levels in H1299^PABPC1–/–
cells re-expressing Myc-PABPC1-WT/K512R under AS treatment at indicated
time points. h Mitophagy flux measured by mtKeima-based flow cytometry
in H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R under AS
treatment. i Live cell confocal imaging of mito-Keima detects
mitophagy. Red mtKeima signal marks mitophagy within lysosomes. Scale
bar, 20 μm. j qPCR quantification of mitochondrial DNA to nuclear DNA
ratio in H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R under
AS treatment. k Mitophagy flux determined by mito-Keima FACS in
H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R with FUNDC1
knockdown under stress condition. CCK-8 assay (l) and plate colony
formation (m) experiments detecting cell viability and clonogenic
ability of H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R with
FUNDC1 knockdown. n Determination of mitophagy flux using mito-keima
FACS assay in H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R,
as well as H1299^PABPC1–/– cells re-expressing Myc-PABPC1-K512R with
FUNDC1 overexpression under stress condition. CCK-8 (o) and plate
colony formation (p) assay evaluating cell viability and clonogenic
ability of H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT/K512R and
H1299^PABPC1–/– cells re-expressing Myc-PABPC1-K512R with FUNDC1
overexpression upon AS treatment. CCK-8 data and plate colony assay are
presented as mean ± SD, n = 6 (a), 4 (b, d, e, l, o) or 5 (c, m, p)
biological replicates, P-values were calculated by two-way ANOVA. qPCR
(f, j) and FACS (P3 cells) (h, k, n) data plotted as mean ± SD of three
biological replicates; P-values determined by one-way ANOVA. Western
blot and Immunofluorescence staining data were representative of at
least 3 biological repeats (g, i).
To comprehensively investigate the role of K512 SUMOylated PABPC1 in
cancer cell survival, we overexpressed Myc-tagged PABPC1-WT or
PABPC1-K512R in both H1299 and H1299^SENP1–/– cells. CCK-8 assays
revealed that cell viability was higher in SENP1 knockout cells
compared to H1299 cells when treated with 20 μM AS for 48 h. Moreover,
overexpression of PABPC1-WT markedly enhanced cell viability relative
to the PABPC1-K512R in both H1299 and H1299^SENP1–/– cells, indicating
the critical role of K512-SUMOylated PABPC1 in promoting cancer cell
survival (Fig. [242]7d). Furthermore, we treated H1299^PABPC1–/– cells
re-expressing either PABPC1-WT or PABPC1-K512R with 20 μM AS and 100 or
150 μM 2-D08, a broad-spectrum SUMOylation inhibitor, for 48 h. CCK-8
assay demonstrated that 2-D08 treatment decreased cell viability in
both PABPC1-WT and PABPC1-K512R cells. Notably, the difference in
viability between PABPC1-WT and K512R mutant cells became insignificant
at 150 µM 2-D08, likely due to the complete inhibition of PABPC1
SUMOylation, which abolished its regulatory function in cell survival
(Fig. [243]7e). Furthermore, in the xenograft model using
HeLa^PABPC1-/- cells re-expressing either Myc-PABPC1-WT or
Myc-PABPC1-K512R, no significant differences in tumor growth or tumor
weight were observed between the PABPC1-WT and PABPC1-K512R groups
under vehicle (PBS) treatment. However, upon treatment with Doxorubicin
(Doxo), a chemotherapeutic agent known to induce cellular stress and
trigger SG formation^[244]9, the PABPC1-K512R group showed a reduction
in tumor growth and tumor weight compared to the PABPC1-WT group
(Supplementary Figs. [245]7c–e). Together, these findings emphasize the
essential role of PABPC1 SUMOylation in supporting cancer cell survival
and growth under stress conditions, with K512 identified as a key SUMO
modification site critical for this regulatory mechanism.
We further explored whether SUMOylated PABPC1 impacts cancer cell
survival under stress by modulating mitophagy. We assessed the mRNA
levels of mitophagy-related genes FUNDC1, BNIP3L, and BECN1. The qPCR
results showed a reduction in these three transcripts in PABPC1-K512R
cells compared to PABPC1-WT cells following AS treatment
(Fig. [246]7f). Subsequent Western blotting analysis provided further
evidence, showing a discernible decrease in the expression levels of
these three proteins in the PABPC1-K512R cells under same stress
condition (Fig. [247]7g, Supplementary Fig. [248]7f). Next, to evaluate
mitophagy flux, we introduced the Mito-keima mitophagy reporter into
H1299^PABPC1–/– and HeLa^PABPC1–/– cell lines stably expressing either
PABPC1-WT or PABPC1-K512R, and mitophagy was quantified by Fluorescence
Activated Cell Sorting (FACS) analysis (gate P3) (Supplementary
Fig. [249]7g). The FACS results showed minimal mitophagy under normal
conditions; however, mitophagy increased upon AS treatment, with
PABPC1-WT cells exhibiting a higher level of mitophagy than
PABPC1-K512R cells (Fig. [250]7h, Supplementary Fig. [251]7h).
Live-cell imaging corroborated these findings, revealing that no red
fluorescence (indicating the absence of mitophagy) was observed under
normal conditions. In contrast, an increase in red fluorescence was
detected in response to AS treatment in H1299^PABPC1–/– cells
re-expressing PABPC1-WT, compared to those re-expressing PABPC1-K512R
(Fig. [252]7i). Moreover, treatment with bafilomycin A1 (Baf-A1), a
well-established autophagy inhibitor, for 24 h nearly abolished red
fluorescence in both PABPC1-WT and PABPC1-K512R cells. This confirmed
that the red fluorescence observed under AS treatment was indeed
indicative of mitophagy and that SUMOylation of PABPC1 at the K512 site
enhanced mitophagy (Supplementary Fig. [253]7i). LC3B is known to
interact with mitophagy receptors through LC3-interacting region (LIR)
motifs to initiate mitophagy in mammalian cells^[254]6.
Immunofluorescence staining showed that PABPC1-WT cells exhibited a
greater number of LC3B foci colocalized with FUNDC1 compared to
PABPC1-K512R cells under stress conditions (Supplementary
Fig. [255]7j), indicating a higher level of mitophagy. Additionally, we
measured the mtDNA/nDNA ratio by qPCR, as a decrease in mtDNA copy
number is a known indicator of mitophagy activation^[256]50. Our
results demonstrated that AS treatment led to a reduction in the
mtDNA/nDNA ratio, with PABPC1-WT cells showing lower ratio compared to
PABPC1-K512R cells, suggesting a higher level of mitophagy in PABPC1-WT
cells (Fig. [257]7j). Collectively, these findings indicate that
SUMOylation of PABPC1 enhances mitophagy during stress.
To further explore the impact of PABPC1 SUMOylation on tumor cell
survival via mitophagy regulation, we silenced the essential mitophagy
gene FUNDC1 in H1299^PABPC1–/– cells re-expressing Myc-PABPC1-WT or
Myc-PABPC1-K512R (Supplementary Fig. [258]7k). Flow cytometry analysis
of mtKeima fluorescence revealed that FUNDC1 knockdown significantly
suppressed mitophagy in H1299^PABPC1-/--Myc-PABPC1-WT cells, whereas no
significant change occurred in mitophagy levels in
H1299^PABPC1-/--Myc-PABPC1-K512R cells (Fig. [259]7k). Viability and
clonogenic survival assays under stress conditions demonstrated that
FUNDC1 knockdown markedly reduced cell viability and colony formation
in H1299^PABPC1–/–-Myc-PABPC1-WT cells, while its impact was minimal in
H1299^PABPC1–/–-Myc-PABPC1-K512R cells (Fig. [260]7l-m). Subsequent
rescue experiments involved overexpressing FUNDC1 in
H1299^PABPC1–/–-Myc-PABPC1-K512R cells (Supplementary Fig. [261]7l),
resulting in substantially restored mitophagy levels under stress
conditions (Fig. [262]7n). Furthermore, evaluations of cell viability
and clonogenic capacity revealed that H1299^PABPC1–/– Myc-PABPC1-K512R
cells exhibited markedly decreased viability and clonogenic potential
relative to H1299 ^PABPC1–/–-Myc-PABPC1-WT cells. However,
overexpressing FUNDC1 into the H1299^PABPC1–/–-Myc-PABPC1-K512R cells
significantly rescued both cell viability and clonogenic capacity
(Fig. [263]7o, p). Similarly, we overexpressed BNIP3L, another critical
mitophagy gene, in H1299^PABPC1–/–-Myc-PABPC1-K512R cells
(Supplementary Fig. [264]7m). The results demonstrated that BNIP3L
overexpression effectively reversed the reduced mitophagy levels
(Supplementary Fig. [265]7n), cell viability (Supplementary
Fig. [266]7o), and clonogenic capacity under stress (Supplementary
Fig. [267]7p). In summary, these findings suggest that PABPC1
SUMOylation enhances mitophagy by regulating the expression of critical
mitophagy genes, such as FUNDC1 and BNIP3L, ultimately promoting tumor
cell survival under stress.
Discussion
SGs play a critical role in adaptive responses of cancer cells,
contributing significantly to their survival^[268]51. However, the
precise regulatory mechanisms underlying SG-mediated enhancement of
tumor cell adaptability remain incompletely understood. In this study,
we demonstrated that SUMOylation of PABPC1, a core component of SGs,
enhances cancer cell survival by promoting mitophagy during cellular
stress. Various stressors, including oxidative stress, heat shock,
osmotic stress, and nutrient starvation, induce SUMOylation at K512 of
PABPC1 (Fig. [269]1e–h, Supplementary Fig. [270]1f–k, Fig. [271]2f,
Supplementary Fig. [272]2k, m). Our findings establish that PABPC1
SUMOylation benefits cell survival under stress (Fig. [273]7a–e,
Supplementary Fig. [274]7a–e) through modulation of mitophagy. Notably,
the K512R mutation of PABPC1 markedly suppresses mitophagy under stress
(Fig. [275]7h–j, Supplementary Fig. [276]7h–i). Silencing the key
mitophagy gene FUNDC1 decreases mitophagy levels and impairs survival
in PABPC1-WT cells (Fig. [277]7k–m). Conversely, overexpressing FUNDC1
or BNIP3L in PABPC1-K512R cells not only restores mitophagy levels but
also substantially enhances their survival capabilities
(Fig. [278]7n–p, Supplementary Fig. [279]7n–p). These findings
emphasize the essential role of mitophagy in stress resilience mediated
by PABPC1 SUMOylation, shedding light on a novel aspect of SG function
in tumor cell adaptability under stress.
SGs play a pivotal role in determining cell fate by modulating gene
expression through the regulation of mRNA translation^[280]10 and
degradation^[281]9,[282]12. Our findings demonstrate that SUMOylation
of PABPC1 precisely controls the stability of specific mRNAs, which
impacts gene expression during stress. Transcriptome-wide analysis
reveals that SUMOylated PABPC1 stabilizes mRNAs under stress conditions
(Fig. [283]4d, e, Supplementary Fig. [284]4a–e). Furthermore,
integrated analysis of RIP-Seq and 4sU-Seq data indicates that
SUMOylated PABPC1 enhances mRNA binding (Fig. [285]5a–e), protecting it
from degradation (Fig. [286]5g, h). Intriguingly, KEGG pathway
enrichment analysis shows a significant enrichment of mRNAs regulated
by SUMOylated PABPC1 within the mitophagy pathway (Fig. [287]4f,
Supplementary Fig. [288]5a–c). Consistently, 4sU-RT-qPCR results
confirm that critical mitophagy-related genes, such as FUNDC1, BNIP3L,
and BECN1, exhibit enhanced stability in PABPC1-WT cells compared to
PABPC1-K512R cells under stress conditions (Fig. [289]4h–j). Notably,
the K512R mutation substantially reduces both mRNA and protein
expression levels of these key mitophagy genes (Fig. [290]7f, g),
emphasizing the essential role of PABPC1 SUMOylation in modulating gene
expression during stress.
Since merely 10% of the bulk mRNAs are present in SGs^[291]12, it is
likely that SG formation specifically regulates certain transcripts.
Previous studies have indicated specific characteristics of mRNAs
stored in SGs, including extended transcript length, AU-rich elements
in the 3’ UTR, and m6A modifications^[292]14,[293]52. Our findings
demonstrate that under stress conditions, transcripts regulated by
SUMOylated PABPC1 exhibit U-rich sequences within their 3’ UTRs
(Fig. [294]6a, Supplementary data [295]5), a feature also observed in
mitophagy-related genes (Supplementary Fig. [296]6i). And SUMOylation
enhanced the interaction between PABPC1 and U-rich mRNAs under stress
condition (Fig. [297]6b). This suggests that SUMOylated PABPC1
specifically modulates mRNAs with U-rich sequence features during
stress.
Given that PABPC1 is traditionally known as a classical mRNA poly(A)
tail-binding protein, questions have arisen regarding its selective
regulation of U-rich mRNAs. Our investigations have highlighted TIA1,
an RNA-binding protein integral to stress granules (SGs), which is
known for specific binding to U-rich sequences^[298]53,[299]54.
Accumulating evidence suggests that TIA1 targets U-rich sequences in
the 3ʹ UTR of mRNAs, potentially participating in RNA stability
regulation and RNA recruitment to SGs. Consequently, we hypothesized
that SUMOylated PABPC1 might regulate U-rich mRNAs through a specific
interaction with TIA1. Co-IP/WB experiments confirmed that TIA1
interacts with SUMOylated PABPC1 via its SUMO-interacting motif (SIM),
facilitating the formation of a PABPC1-SUMO-TIA1 complex
(Fig. [300]6c–e). Furthermore, TIA1 directly binds to U-rich mRNAs and
co-localizes with PABPC1 in SGs under stress conditions (Supplementary
Fig. [301]6c, Supplementary Fig. [302]6e). RIP-qPCR results also
indicate that stress conditions enhance the interaction between TIA1
and the mRNAs of FUNDC1, BNIP3L, and BECN1 (Fig. [303]6f). In addition,
in vitro poly(U) RNA binding assays indicated that SUMO modification
enhances the binding of PABPC1 to both TIA1 and poly(U) RNA
(Fig. [304]6g). These findings elucidate a targeted mechanism by which
SUMOylated PABPC1 recruits U-rich mRNAs into SGs through the
interaction of attached SUMO1 with TIA1, safeguarding these mRNAs from
degradation during cellular stress responses. This mechanism enhances
our comprehension of the nuanced roles of SG components in mRNA
dynamics.
In summary, our study elucidates a molecular mechanism by which
SUMOylation of PABPC1 regulates the function of SGs and supports cancer
cell survival under stress conditions (Fig. [305]8). Specifically,
stress triggers extensive SUMOylation of PABPC1, leading to
interactions between its covalently attached SUMO1 and the
SUMO-interacting motif (SIM) of TIA1 protein. This interaction fosters
the formation of the PABPC1-SUMO-TIA1 complex, which colocalizes within
SGs. TIA1, within this complex, binds specifically to mRNAs
characterized by U-rich sequences, effectively recruiting these mRNAs
to SGs for protection against degradation. Crucially, this mechanism
extends protection to key mRNAs within the mitophagy pathway, such as
FUNDC1, which feature U-rich sequences in their 3’ UTR. The expression
of these genes under stress conditions facilitates mitophagy, promoting
the clearance of damaged mitochondria and thus enhancing cellular
resilience. This adaptive response is pivotal in maintaining cellular
homeostasis and enhancing the stress adaptability of cancer cells,
ultimately increasing their survival in adverse conditions.
Fig. 8. Model illustrating the role of SUMOylated PABPC1 in stress tolerance.
[306]Fig. 8
[307]Open in a new tab
Under stress conditions (e.g., sodium arsenite, heat shock, osmotic
stress, glucose starvation), PABPC1 undergoes SUMOylation and interacts
with TIA1 to form a PABPC1–SUMO–TIA1 complex. This complex recruits
U-rich mRNAs into stress granules, preventing their degradation and
promoting the expression of mitophagy-related genes such as FUNDC1,
thereby maintaining cellular homeostasis and enhancing cancer cell
stress adaptation.
Limitations of the study
Several questions remain at this stage. First, although we observed
that various stress induce SUMOylation of PABPC1, the molecular
mechanisms regulating SUMOylation of PABPC1 remain to be elucidated.
SUMOylation is catalyzed by the dimeric E1 enzyme SAE1/UBA2, the single
E2 enzyme Ubc9, and E3 ligases, and can be reversed by Sentrin-specific
proteases (SENPs), with the regulation of SUMO levels often controlled
by the activity of E3 ligases and SENP enzymes. The elevation in
SUMOylation levels of PABPC1 under stress could be mediated by enhanced
interactions with a specific E3 ligase or due to a decrease in SENP
enzyme activity under stress conditions. Additionally, mass
spectrometry data indicate that stress granules (SGs) contain
SUMO-related E2 and E3 enzymes, as well as SUMO molecules. The entry of
PABPC1 into SGs under stress brings it into closer spatial proximity to
SUMO-related E2 and E3 enzymes and SUMO molecules, which may also
contribute to the increased levels of SUMOylation.
Based on high-throughput sequencing results, we have observed that
under stress conditions, SUMOylation of PABPC1 specifically regulates
U-rich mRNAs. However, notable stability variations also manifest in
transcripts that lack U-rich sequences between PABPC1-WT and
PABPC1-K512R cells. This leads us to question whether transcripts
regulated by SUMOylated PABPC1 under stress conditions might exhibit
additional sequence features or specific RNA modifications. Elucidating
the presence of these features and their regulatory mechanisms under
stress conditions is a compelling direction for further research.
Methods
Ethics statement
All animal experiments were conducted in accordance with the Guide for
the Care and Use of Laboratory Animals and were approved by the
Institutional Animal Care and Use Committee of Shanghai Jiao Tong
University School of Medicine (IACUC approval NO. JUMC2024-218-A).
Antibodies and reagents
Antibodies against PABP (Ab21060) and SENP1 (ab108981) were purchased
from Abcam. Antibodies against SUMO-1 (4930), His-tag (2366), Myc-tag
(2276), FUNDC1 (49240), and LC3B (83506) were purchased from Cell
Signaling Technology. Antibodies against TIA1 (12133-2-AP), GAPDH
(HRP-60004), Alpha Tubulin (66031-1-Ig), Beta Actin (66009-1-Ig),
BNIP3L (12986-1-AP), and Beclin1 (11306-1-AP) were purchased from
Proteintech. Monoclonal anti-Flag M2 antibody (F1804) was purchased
from Sigma-Aldrich. Monoclonal anti-HA antibody (MMS-101R) was from
Covance. Mouse anti-GST antibody (CW0084) was purchased from CWBio.
Antibodies against G3BP1 (H-10) (sc-365338), Mouse anti-normal mouse
IgG antibody (sc-2025), and Rabbit anti-normal rabbit IgG (sc-2027)
were purchased from Santa Cruz Biotechnology.
Plasmids
Human PABPC1, TIA1, FUNDC1, BNIP3L genes were amplified by KOD-plus Kit
(TOYOBO) using human cDNA made from HEK293T cell and then subsequently
cloned into pCMV-HA, pCMV-Myc, pGEX-4T-1, or lentivector-based pCD510B
vectors, respectively. Point mutated PABPC1 and SIM mutated TIA1 were
introduced by using KOD-plus-mutagenesis Kit (TOYOBO) according to the
manufacturer’s protocol. PABPC1, FUNDC1 shRNA oligoes were subcloned
into the lentiviral vector pLKO.1. The sgRNAs against PABPC1 were
designed according to the previous report^[308]55 then cloned into
lentiCRISPRv2 (Addgene) via DNA Ligation Kit (Vazyme). The plasmids
were extracted using NucleoBond Xtra Midi Plus kit (Macherey-Nagel) and
validated by Sanger sequencing (Sangon Biotech). The plasmid
pHAGE-mt-mKeima was purchased from Addgene
([309]http://n2t.net/addgene:131626; RRID: Addgene_131626). All the
primers and oligoes used for molecular cloning were listed in
Supplementary data [310]1.
Cell line and cell culture
The human HEK-293T, HEK-293FT, H1299, and HeLa cell lines were obtained
from National Collection of Authenticated Cell Cultures, Shanghai,
China. All these cell lines were cultured in DMEM medium (Corning)
supplemented with 10% fetal bovine serum (Yeasen) and 1%
penicillin/streptomycin (Yeasen) at 37 °C in a 5% CO[2] humidified
incubator. The culture medium was changed 24 h before stress
conditions. Arsenite stress was induced by treating with 0.5 mM AS
(sodium arsenite, Sigma-Aldrich) for 1–3 h as indicated, and osmotic
stress was triggered using 0.4 M sorbitol for 0.5-4 h. For heat shock
experiments, cells were incubated at 46 °C for 1 h. For glucose
starvation experiments, DMEM medium were replaced by DMEM without
L-glutamine medium for indicated time.
Cell transfection and lentivirus infection
Cell transfections were performed by using PEI for HEK-293T and
HEK-293FT, and Lipofectamine 2000 for HeLa and H1299 cells following
the manufacturer’s protocol. To infect H1299 and HeLa cells with
pseudo-lentivirus, cells were digested with Trypsin-EDTA (Yeasen) to
single cells and cultured in corresponding medium with lentivirus and
10 mg/mL polybrene for 48 h. Infected cells were cultured for one more
day with the addition of 2-10 μg/ml puromycin for at least 2 days.
After that, proteins were collected for further detection.
SUMOylation assays
SUMOylation of PABPC1 was confirm by three different methods in vivo
and in vitro. The method of SUMOylation assay through Ni^2+-NTA
pulldown was performed as described^[311]29,[312]31,[313]56–[314]59.
The detection of both exogenous and endogenous SUMOylation of PABPC1
was followed by co-immunoprecipitation (Co-IP) under denaturing
conditions as published protocol^[315]60,[316]61, with minor changes as
our previous publication described^[317]30,[318]59. In brief, cells
were lysed in denature lysis buffer I (50 mM Tris-HCl, pH 6.8, 2% SDS,
40 mM DTT, and 5% glycerol), boiled for 10 min, and then sonicated
until the lysate became fluid. The lysate was cleared by centrifuge at
Room temperature (17,000 × g, 10 min) and diluted 1:10 with cold
denature lysis buffer II (20 mM NEM, 50 mM Tris-HCl, pH 7.4, 150 mM
NaCl and 1% Nonidet P-40). Finally, Protein A/G beads and antibody
(anti-PABPC1) were added into the lysates. After overnight of rotation
at 4 °C, the beads were washed five times in cold denature lysis buffer
II before SDS-PAGE loading buffer was added to the beads.
The method of in vitro E.coli BL21-based SUMOylation assay was
performed as previous described^[319]58,[320]59. Briefly, the
GST-tagged recombinant PABPC1 plasmid was transformed to BL21 (DE3)
Escherichia coli cells alone or with pT-E1E2S1 plasmid, the bacterial
clone that express both PABPC1 and pE1E2S1 proteins were then induced
at 16 °C with 0.5 mM IPTG for 16–20 h. Bacteria were collected and
lysed in PBS-L buffer (50 mM Tris-HCl pH7.4, 150 mM NaCl) and sonicated
for 10 min on ice. The lysate was then cleared by centrifuge at 4 °C
(17, 000 g, 30 min). The proteins were affinity purified using
GST-Sefinose Resin. After extensive wash with PBS-L buffer, proteins
were eluted with GSH buffer (50 mM Tris pH 8.0, 20 mM GSH). The
purified protein was verified by Western blotting.
Western blotting
Cell lysates or protein solutions were mixed with 6× protein loading
buffer (Sangon Biotech) and boiled for 10 min. Following rapid
centrifugation, the protein samples were loaded onto an SDS-PAGE gel
for electrophoresis. The separated proteins were subsequently
transferred onto the 0.22 μm PVDF membrane and blocked at room
temperature with 5% milk in TBS with 0.1% Tween 20 (TBST) for 1 h.
Then, the membrane was incubated with primary antibody overnight at
4 °C, washed 3 times for 5 min each with TBST, incubated with secondary
antibody for 1 h at room temperature, washed 3 times of 5 min with
TBST, and then incubated with the ECL dection buffers (Tanon). The
protein signals were detected via Amersham Imager (GE HealthCare).
Co-Immunoprecipitation (Co-IP)
For Co-IP analysis, HEK-293T or H1299 cells were transfected with
protein expression plasmids for 48 h or treated with AS for indicated
times. Then, cells were harvested and suspended in 1 mL RIPA lysis
buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 40 mM N-Ethylmaleimide, 1%
NP-40, protease inhibitor cocktail) followed by sonication for 10 times
of 3 sec. After 30 min of 17,000 × g centrifugation at 4 °C, the
supernatant was collected as protein lysates. 1 mg total protein were
incubated with with 20 μL of protein A/G beads (Santa Cruz) and 1 μg of
primary antibody overnight at 4 °C. The beads were then washed 5 ×
5 min with RIPA buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 40 mM
N-Ethylmaleimide, 1% NP-40) and mixed with 2× protein loading buffer.
After boiling for 10 min, the protein sample were further analyzed by
western blotting.
Oligo(dT) pulldown
For Oligo(dT) pulldown experiment, HEK-293T cells were transfected with
protein expression plasmids for 48 h. Cells were lysed with lysis
buffer (20 mM Tris-HCl pH 7.4, 250 mM NaCl, 10 mM KCl, 5 mM MgCl[2],
0.1% Triton-X, 40 mM N-Ethylmaleimide, RNase inhibitor, protease
inhibitors) on ice for 1 h followed by 10× 3 s sonication. After 20 min
of 13,000 × g centrifugation at 4 °C, the supernatant was collected as
protein lysates. 20 μL of Dynabeads™ Oligo(dT)[25] were then incubated
with 1 mg cell lysate overnight at 4 °C. Finally, the beads were washed
with RIP buffer (20 mM Tris-HCl pH 7.4, 250 mM NaCl, 10 mM KCl, 5 mM
MgCl[2], 0.1% Triton-X, 40 mM N-Ethylmaleimide) for 5 times, and the
proteins were detected by western blotting.
Immunofluorescence staining and microscopy
For immunofluorescence (IF) staining, cells cultured on glass
coverslips in 24-well plates were treated with AS at different doses
for indicated time and then washed by PBS. Cells were fixed using 4%
paraformaldehyde in PBS for 15 min at room temperature and
permeabilized with PBS containing 0.5% Triton X-100 at room temperature
for 60 min. After blocking with 5% bovine serum albumin in PBST
(PBS + 0.5% Tween 20) for 60 min, SGs were immunostained with rabbit
anti-PABPC1 or mouse anti-G3BP1 overnight at 4 °C. 488 goat anti-rabbit
IgG antibody and 568 goat anti-mouse IgG antibody were diluted in
PBS-BSA supplemented with 2 μg/ml of Hoechst and stain at RT for
60 min. Cells were washed 3 times with PBST and then mounted on glass
slides in Prolong Gold mounting agent. Fluorescence images were
collected using Leica TCS SP8 or Leica TCS Sp8 STED confocal
microscope. The number of stress granules foci were counted with
CellProFiler software^[321]62.
Fluorescence recovery after photobleaching (FRAP) assay
In each FRAP experiment, three regions of interest (ROIs) of identical
size were selected. ROI1 was the area where photobleaching was applied,
ROI2 served as a comparable droplet for correcting system fluorescence
fluctuations, and ROI3 was located in the surrounding dilute solution
for background correction. A 0.8 μm diameter circle at the center of a
droplet was selected for bleaching. After capturing an initial image, a
488 nm laser was used to bleach ROI1 until the fluorescence intensity
reached a level similar to that of ROI3. Images were captured by Zeiss
900 and analyzed using ZEN v3.8, with the pre-bleach fluorescence
intensity set to 1. Data are presented as mean ± SD.
Isolation of PABPC1 interaction SG proteins for LC-MS/MS
For identification of the proteome profile associated by PABPC1 in
stress granule, H1299 cells stable expressing Myc-PABPC1 were grown to
85% confluency in two 10 cm^2 square culture dishes and treated with
0.5 mM AS for 1 h at 37 °C/5% CO[2]. For the recovery group, cells
treated with AS recovered from stress by replacing the normal media.
After treatment, cells were cross-linked using fresh formadehyde (1%)
in culture media for 10 min at room temperature, and quenched with
glycine at a final concentration of 125 mM. Then cells were washed once
with PBS containing 0.1%NP-40, transferred to falcon tubes, and
pelleted at 800 g for 3 min. Upon aspirating the media, the pellets
were immediately flash-frozen in liquid N[2] and stored at -80 °C until
isolation of SG cores was performed.
The isolation of SG cores was adapted from three
papers^[322]12,[323]21,[324]63. Briefly, the pellet was re-suspended in
SG lysis buffer (50 mM Tris-HCl pH7.4, 100 mM KOAc, 2 mM MgOAc, 0.5 mM
DTT, 0.5%NP-40, 40 mM NEMI, and Protease inhibitor cocktail) and
sonicated for 20 s on ice. After lysis, the lysates were spun at 1,
000 g for 5 min at 4°C to pellet cell debris and extracted SG core as
following steps: (1) The supernatant was transferred to new tube then
spun at 17,000 × g for 30 min at 4 °C to pellet SG cores. (2) The
resulting supernatant was discarded and the pellet was re-suspended in
1 mL of SG lysis buffer. (3) Step1 and 2 were repeated to enrich for
SG. (4) The resulting pellet was washed twice with SG lysis buffer and
re-suspended in 1 mL of SG lysis buffer, then sonicated for 10 s on ice
and spun at 850 g for 2 min at 4 °C. (5) The supernatant which
represents the SG core enriched fraction was transferred to a new tube
and pre-cleared by adding 10 μL of protein A/G beads and nutating at
4 °C for 1 h then beads were removed. (6) 2 μL of anti-Myc antibody and
30 μL of protein A/G beads which pre-blocked with 1% BSA were added to
the enriched fraction and incubated at 4°C overnight to affinity purify
SG cores. (7) Beads were then washed three times for 5 min each in
buffer1 (20 mM Tris-HCl pH 7.4, 200 mM NaCl, 0.01% NP40), 5 min in
buffer2 (20 mM Tris-HCl pH 8.0, 500 mM NaCl, 0.01% NP40), and two times
for 5 min each in buffer3 (20 mM Tris-HCl pH 8.0, 200 mM NaCl, 0.01%
NP-40, 0.5 mM DTT, 5 mM EDTA). (8) Following the final wash, beads were
resuspended in 2% SDS and boiled at 100 °C for 10 min. Supernatant was
performed for Mass Spectrometric analysis.
Mass spectrometry (MS) analysis
For Mass spectrometry (MS) analysis, samples were prepared using the
PABPC1 interaction SG proteins isolation protocol described above, with
two biological replicates included per group. Mass spectrometric
experiments were performed on Orbitrap Fusion LUMOS mass spectrometer
(Thermo Fisher Scientific) coupled to an Easy-nLC 1200 via an Easy
Spray (Thermo Fisher Scientific). The peptides mixtures were loaded to
the reverse-phase microcapillary column (0.1 × 150 mm) packed with
Reversed Phase C18 resins (2 μm, PepMap RSLC) at a flow of 1 μl/min and
separated using a 60-min linear gradient solution from 95% buffer A
(0.1% formic acid, 2% acetonitrile and 98% water) to 30% buffer B (0.1%
formic acid and 80% acetonitrile) at a flow rate of 0.3 μl/min. The
spray voltage was set to 2.1 KV, with the temperature of the ion
transfer capillary set at 275°C, and the radio frequency (RF) lens was
60%. The mass spectrometer was operated in positive ion mode and
employed in the data-dependent mode to automatically switch between
mass spectrometry and tandem mass spectrometry (MS/MS) within the
specialized cycle time (3 s).
One full mass spectrometry scan from 350-1, 500 m/z was acquired at
high resolution R = 120, 000 (defined at m/z = 400) and MS/MS scans
were acquired at a resolution of 30,000. Masses selected for MS/MS were
isolated (quadrupole) at a width of 4 Da and were fragmented using a
higher energy collisional dissociation of 30% ± 5. All MS/MS ion
spectra were analyzed using PEAKS 10.0 (Bioinformatics Solutions) for
processing, de novo sequencing and database searching. The resulting
sequences were searched against the UniProtHuman Proteome database. FDR
estimation was enabled. Peptides were filtered for −log10(P value) ≥20,
and proteins were filtered for −log10(P value) ≥ 15 plus one unique
peptide. For all of the experiments, these setting gave an FDR < 1% at
the peptide-spectrum match level. Proteins sharing significant peptide
evidence were grouped into clusters. The relative ratios (PABPC1-K512R/
PABPC1-WT) of proteins associated with PABPC1 under sodium arsenite
treatment and recovery were calculated by normalizing to PABPC1 itself.
The mass spectrometry (MS)-based proteomics was performed at the
Proteomics of Core Facility of Basic Medical Sciences, Shanghai Jiao
Tong University School of Medicine (SJTU-SM). The mass spectrometry
data were analyzed using Perseus_v2.1.3.0 software and visualized using
GraphPad Prism 8 software.
mRNA stability profiling
mRNA stability profiling was performed using 4-Thiouridine (4sU)
Pulse-Chase sequencing (4sU-Seq) method. Myc-PABPC1 stably expression
cells were labeled with 200 μM 4-thio-uridine (Sigma-Aldrich) for 2 h,
after which medium was replaced with fresh medium lacking
4-thio-uridine. Cells were harvested after an additional 0, 3, 6 h.
Total cell RNA was isolated using TRIzol Reagent (Sigma-AldRich).
Subsequently, 4-thio-uridine-labeled RNA was isolated following a
manufacturer’s instruction of Dölken^[325]64. Briefly,
4-thio-uridine-containing RNAs were converted to biotinylated RNAs
using HPDP-Biotin (Thermo Fisher Scientific) in N, N-dimethylformamide.
The resulting biotinylated RNAs were isolated using Dynabeads^TM
MyOne^TM Streptavidin C1 (Thermo Fisher Scientific). Isolated RNAs were
subjected to RNA-seq or RT-qPCR (4sU-RT-qPCR). For 4sU-RT-qPCR, mRNA
levels were detected by mRNA-specific primers and normalized to the
level of ACTB mRNA.
The mRNA stability was also assessed by qPCR in actinomycin D
(ActD)-treated cells. Briefly, cells were treated with 5 μg/mL ActD and
collected at the specified time points. Total RNA was extracted using
Trizol and analyzed by qPCR, with ACTB mRNA used as the normalization
control. The mRNA stability was then quantified using GraphPad Prism 8.
RNA immunoprecipitation assay (RIP)
RIP was performed as our previous study^[326]65 with some modification.
Briefly, stable cell lines seeded in a 10-cm Dish at 80–90% confluency
were treated with or without AS and harvested by Trypsinization. Cells
were lysed in RIP-lysis buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl,
10 mM EDTA, 2 mM MgCl[2], 1 mM DTT, 0.5% NP-40, 100 units/ml RNase
inhibitor, 400 μM VRC and Protease inhibitor cocktail) on ice for 1 h.
1/10 of cell lysates were extracted with 1 mL of TRIzol reagent
(Sigma-AldRich) for extraction of total RNAs as input, 1/50 of cell
lysates were saved for Western blotting to detect the protein
expression, and left lysates were incubated with indicated antibodies
and protein A/G-agarose beads at 4 °C for 3 h. After washing with
RIP-lysis buffer for five times, 1/10 of beads was subjected to Western
blotting analysis to identify the efficiency of IP, and the remained
beads were extracted with 1 mL of TRIzol reagent for extraction of RIP
bound RNAs. Input and co-immunoprecipitated RNAs were analyzed by
RT-qPCR or RNA-Seq.
RT-qPCR
RNAs were extracted by TRIzol reagent (Sigma-AldRich) and then treated
with DNase I (Thermo Fish Scientific) to degrade genomic DNA. The
reverse transcription was performed with 1 μg total RNA and the
PrimeScript^TM RT Reagent Kit (Takara) according to the manufacturer’s
instructions. Quantitative real-time PCR (RT-qPCR) was conducted using
SYBR Green PCR Master Mix (Yeasen) to determine the mRNA expression
level of a gene of interest. Gene expression levels were normalized to
the expression of ACTB.
High-throughput sequencing for 4sU-Seq and RIP-Seq
For 4sU-Seq, 4sU-labled RNA were extracted as described above for
removing the rRNAs using Ribo-Zero rRNA Removal Kits (Illumina, San
Diego, CA, USA) as the manufacturer’s instructions for the following
library construction. The rRNA-depleted RNAs were constructed RNA
sequencing libraries by using TruSeq Stranded Total RNA Library Prep
Kit (Illumina, San Diego, CA, USA) according to the manufacturer’s
instructions.
For RIP-Seq, the indicated stable H1299 cell lines were treated with or
without AS, and RNA was immunoprecipitated as described above. The
immunoprecipitated RNA was then processed for sequencing according to
the outlined protocol^[327]66. In brief, RNAs bound to PABPC1 and total
RNAs (as an input) were extracted by using TRIZOL reagent as following
manufacturer’s instruction (Invitrogen), then the rRNAs were removed
from the immunoprecipitated RNA and input RNA samples by using RNAs
with NEBNext rRNA Depletion Kit (New England Biolabs, Inc.,
Massachusetts, USA). The rRNA-depleted RNAs were constructed RNA
sequencing libraries by using NEBNext® Ultra™ II Directional RNA
Library Prep Kit (New England Biolabs, Inc., Massachusetts, USA)
according to the manufacturer’s instructions. Constructed 4sU-Seq and
RIP-Seq libraries were controlled for quality and quantified using the
BioAnalyzer 2100 system (Agilent Technologies, Inc., USA), and the
libraries sequencing were performed on an illumina Hiseq instrument
with 150 bp paired-end reads.
High-throughput sequencings for 4sU-Seq and RIP-Seq were all done by
Cloud-Seq Biotech (Shanghai, China).
Analysis for high-throughput sequencing data
For 4sU-Seq data, samples were sequenced by Illumina NovaSeq 6000
platform. After sequencing, reads were trimmed for adaptor sequence,
masked for low-quality sequence by Cutadapt^[328]67 and then mapped to
human reference genome (UCSC hg19) using Hisat2^[329]68. Library depth
normalized gene expression counts were calculated using DESeq2^[330]69.
Then, all normalized gene count values in a particular time point were
divided by the probe value for that time point. Data was further
normalized to the first time point and log transformed. Half-lives were
calculated using BridgeR2 (version 0.1.0)^[331]70,[332]71. A maximum
half-life value of 24 hrs was applied to calculated values exceeding
this value. Genes with R^2 values of <0.8 were excluded from further
analysis. Only genes with protein coding annotation were used for
downstream analysis.
For RIP-seq data, samples were sequenced by Illumina NovaSeq 6000
platform. After sequencing, all reads were mapped tohuman reference
genome (UCSC hg19) by Hisat2 software with default parameters. The
target binding regions of PABPC1 were identified using MACS
software^[333]72. High-confidence binding regions of PABPC1 were
identified by stringent cutoff threshold, and then annotated with the
latest UCSC RefSeq database to connect the peak information with the
gene annotation. Fold enrichment of each mRNA transcript matched to
different MACS identified motifs were summed.
For Sequence motif analysis, The RNA sequence from 4sU-seq and RIP-seq
were analyzed for the occurrence of over-represented motifs. We
performed de-novo motif finding using MEME (version 5.5.5) in
standalone mode. We ran MEME with a maximum motif width of 20 bp, and
also with unrestricted length. Sequence logos presented were produced
within MEME output.
For pathway enrichment analysis, the GO and KEGG pathway enrichment
analysis was performed using David bioinformatics resources.
Electrophoretic mobility shift assay (EMSA)
50 nM of Biotin-labeled RNAs were used for each electrophoretic
mobility shift assay (EMSA) reaction. Protein-RNA incubation was
carried out with the indicated amount of purified protein and
Biotin-labeled RNA in EMSA buffer (125 mM Tris-HCl pH 7.5, 1.25 M KCl,
25 mM MgCl[2], 50% Glycerol, 10 mg/mL BSA, 1 unite RNase Inhibitor) at
room temperature for 30 min. The binding reactions were then mixed with
5× EMSA loading buffer (75% Glycerol, 2.5× TBE, 0.06% bromophenol blue,
0.06% xylene cyanol) and loaded onto a 7% native page gel for
electrophoresis. After that, the separated protein-RNA were transferred
onto the Nylon membrane (Sigma-Aldrich) fixed to the membrane by UV
crosslinking (480 mJ/cm^2). The membrane was then incubated with
Stabilized streptavidin-HRP Conjugate (Thermo Fish Scientific) at room
temperature for 1 h, washed 3 times of 10 min with 1× Washing buffer
(Thermo Fish Scientific). The signal of Biotin-labeled RNA were
detected by Amersham Imager (GE HealthCare).
In vitro poly(U) RNA binding assay
For the in vitro poly(U) RNA binding assay, protein-RNA incubation was
performed with 2 μM purified His-GFP-PABPC1 or
His-GFP-PABPC1 + pE1E2S1, 2 μM GST-TIA1 protein, and 50 nM
biotin-labeled poly(U) RNA in reaction buffer (20 mM Tris-HCl pH 7.5,
200 mM KCl, 2 mM MgCl₂, 5% glycerol, 1 mM DTT, 40 U RNase inhibitor)
overnight at 4 °C. The resulting protein complexes were then captured
using Ni²⁺-NTA resin precipitation to pull down PABPC1. After washing
away unbound protein and poly(U) RNA with RIP-lysis buffer, the RNA was
purified from the beads and analyzed by northern blotting, while the
associated proteins were detected by Western blotting.
Mito-Keima mitophagy assay
To monitor mitophagy, we used mito-keima (mitochondria-targeted Keima),
a ratiometric, pH-sensitive fluorescent protein that is resistant to
lysosomal proteases^[334]73. As Keima is characteristically excited at
561 nm under acidic conditions and at 488 nm under neutral pH
environments, a high ratio of 561/488 nm fluorescence represents the
presence of mitochondria in acidic lysosomes (mitochondria actively
undergoing mitophagy). For quantitation of mitophagy by FACS, cells
stably expressing mito-keima following arsenite treatment were
harvested by trypsinization and resuspended in FACS buffer (145 mM
NaCl, 5 mM KCl, 1.8 mM CaCl[2], 0.8 mM MgCl[2], 10 mM HEPES, 10 mM
glucose, 0.1% BSA). Measurements of lysosomal Mitokeima were made using
dual-excitation ratiometric pH measurements at 488 (pH 7) and 561 (pH
4) nm lasers with 620/29 nm and 614/20 nm emission filters,
respectively. For each sample, 10,000 events were collected. Data were
analyzed using FlowJo. In addition, cells stably expressing mito-Keima
could be imaged by Leica confocal microscope.
mtDNA copy number
The mitochondrial DNA (mtDNA) content relative to nuclear DNA (nDNA)
was determined as previously described^[335]50. Briefly, cells treated
with or without AS were collected, and total DNA was extracted using a
DNA Extraction Kit (Biosharp). qPCR was then performed to measure mtDNA
(tRNA-Leu (UUR)) and nDNA (B2m/β2-microglobulin) levels. The relative
mtDNA/nDNA ratio was calculated using the following equations:
[MATH: △Ct=(nDNACt−mtDNACt) :MATH]
1
[MATH: RelativemitochondrialDNAcontent=2×2ΔCt :MATH]
2
Cell proliferation assay
Cell proliferation was assessed using the Cell Counting Kit-8 (CCK-8)
(Yeasen), a highly sensitive and widely used assay for evaluating cell
proliferation and cytotoxicity, based on the WST-8 reagent (chemical
name:
2-(2-methoxy-4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfonatophenyl)
-2H-tetrazolium monosodium salt). In this assay, WST-8 is reduced by
mitochondrial dehydrogenases in the presence of electron-coupling
reagents, resulting in the formation of a soluble orange-yellow
formazan product. The intensity of the resulting color is directly
proportional to the number of proliferating cells and inversely
proportional to cytotoxicity. The optical density (OD) at 450 nm was
measured using a microplate reader, providing an indirect assessment of
viable cell count.
For the proliferation assay, H1299 and HeLa cells were infected with
the indicated lentivirus and selected for 1 week to generate stable
knockout or overexpression cell lines. The cells were seeded into
96-well plates at a density of 1 × 10^3 cells per well, in a final
volume of 100 μL. At each designated time point, 10 μL of CCK-8
solution was added to each well, and the absorbance at 450 nm was
measured to evaluate cell proliferation.
Cell viability and Clonogenic survival assay
For cell viability assay, cells were counted and 1 × 10^4 cells were
seeded into 96 well plates and allowed to adhere for 24 h before being
treated with AS for 48 h. Viable cells was determined using CCK8-kit
and all quantitative results were normalized to non-treatment group.
For clonogenic survival assay, 2 × 10^3 cells were seeded into a
12-well plate and cultured for 3 days. AS was added into medium and all
culture medium was changed every 3 days until colony was visible (7-14
days). Colonies were washed, fixed and stained with 0.1% (w/v) crystal
violet overnight. Visible colonies were counted and analyzed between
groups with ImageJ.
Xenograft tumor model
All animal experiments were conducted in accordance with the Guide for
the Care and Use of Laboratory Animals and approved by the
Institutional Animal Care and Use Committee of Shanghai Jiao Tong
University School of Medicine (Approval NO. JUMC2024-218-A). Mice were
obtained from Shanghai Lingchang Biotechnology Co., Ltd. (Shanghai,
China), housed in a specific pathogen-free environment, and allowed to
acclimate to the conditions under careful handling prior to
experimentation.
The xenograft tumor model was established as previously described.
Briefly, HeLa^PABPC1-/- cells stably expressing Myc-PABPC1-WT/K512R
(4×10^6) were subcutaneously injected into 7-week-old male BALB/c nude
mice. Mice were maintained on a standard chow diet (Xietong Shengwu,
catalog no. XTC01WC-001) ad libitum under specific pathogen-free (SPF)
conditions. Mice were housed in the animal facility of Shanghai Jiao
Tong University School of Medicine under a 12-h light/dark cycle, at a
controlled temperature of 20–22 °C and 60% relative humidity. Once the
tumors reached approximately 100 mm³, the mice were treated with
doxorubicin (MedChemExpress) via intraperitoneal (i.p.) injection at a
dose of 1 mg/kg every three days for a total of four injections. The
control group received an equivalent volume of vehicle (PBS) via
intraperitoneal injection. Tumor growth was monitored with calipers,
and tumor volume was calculated using the formula: (length × width² /
2). Mice were euthanized when they met the institutional criteria for
tumor size (2 cm). At the study endpoint, tumors were harvested and
weighed. All animal studies were conducted with the approval and
guidance of the Shanghai Jiao Tong University Medical Animal Ethics
Committees.
Statistical analysis
Statistics in this study were presented as mean ± SD. Error bars
represented SD in triplicate experiments if not mentioned otherwise.
Statistical comparisons were performed by using two-tailed t tests,
one-way ANOVA, two-way ANOVA, or twosided Mann–Whitney U test as
indicated in the figure legends. Statistical analyses were carried out
using Graphpad Prism 8 (GraphPad Software). Each sequence RNA sample
(4sU-seq and RIP-seq) has two biological replicates. For other
experiments, the number of replicates is indicated in the figure
legends.
Reporting summary
Further information on research design is available in the [336]Nature
Portfolio Reporting Summary linked to this article.
Supplementary information
[337]Supplementary Information^ (6.5MB, pdf)
[338]41467_2025_62619_MOESM2_ESM.pdf^ (234.3KB, pdf)
Description of Additional Supplementary Files
[339]Supplementary Data 1^ (15.2KB, xlsx)
[340]Supplementary Data 2^ (1.6MB, xlsx)
[341]Supplementary Data 3^ (16.8MB, xlsx)
[342]Supplementary Data 4^ (9.1MB, xlsx)
[343]Supplementary Data 5^ (213.2KB, xlsx)
[344]Reporting Summary^ (190.8KB, pdf)
[345]Transparent Peer Review file^ (924.4KB, pdf)
Source data
[346]Source Data^ (26.7MB, xlsx)
Acknowledgements