Abstract
Background
Increasing evidence supports the role of small nucleolar RNAs (snoRNAs)
and long non-coding RNAs (lncRNAs) as master gene regulators at the
epigenetic modification level. However, the underlying mechanism of
these functional ncRNAs in colorectal cancer (CRC) has not been well
investigated.
Methods
The dysregulated expression profiling of lncRNAs-snoRNAs-mRNAs and
their correlations and co-expression enrichment were assessed by
GeneChip microarray analysis. The candidate lncRNAs, snoRNAs, and
target genes were detected by in situ hybridization (ISH), RT-PCR, qPCR
and immunofluorescence (IF) assays. The biological functions of these
factors were investigated using in vitro and in vivo studies that
included CCK8, trans-well, cell apoptosis, IF assay, western blot
method, and the xenograft mice models. rRNA 2′-O-methylation (Me)
activities were determined by the RTL-P assay and a novel
double-stranded primer based on the single-stranded toehold (DPBST)
assay. The underlying molecular mechanisms were explored by
bioinformatics and RNA stability, RNA fluorescence ISH, RNA pull-down
and translation inhibition assays.
Results
To demonstrate the involvement of lncRNA and snoRNAs in 2′-O-Me
modification during tumorigenesis, we uncovered a previously unreported
mechanism linking the snoRNPs NOP58 regulated by ZFAS1 in control of
SNORD12C, SNORD78 mediated rRNA 2′-O-Me activities in CRC initiation
and development. Specifically, ZFAS1 exerts its oncogenic functions and
significantly up-regulated accompanied by elevated NOP58, SNORD12C/78
expression in CRC cells and tissues. ZFAS1 knockdown suppressed CRC
cell proliferation, migration, and increased cell apoptosis, and this
inhibitory effect could be reversed by NOP58 overexpression in vitro
and in vivo. Mechanistically, the NOP58 protein could be recognized by
the specific motif (AAGA or CAGA) of ZFAS1. This event accelerates the
assembly of SNORD12C/78 to allow for further guiding of 2′-O-Me at the
corresponding Gm3878 and Gm4593 sites. Importantly, silencing SNORD12C
or 78 reduced the rRNAs 2′-O-Me activities, which could be rescued by
overexpression ZFAS1, and this subsequently inhibits the RNA stability
and translation activity of their downstream targets (e.g., EIF4A3 and
LAMC2).
Conclusion
The novel ZFAS1-NOP58-SNORD12C/78-EIF4A3/LAMC2 signaling axis that
functions in CRC tumorigenesis provides a better understanding
regarding the role of lncRNA-snoRNP-mediated rRNAs 2′-O-Me activities
for the prevention and treatment of CRC.
Keywords: ZFAS1, SNORD12C, SNORD78, 2′-O-methylation (2′-O-me), NOP58,
Colorectal cancer (CRC)
Introduction
Recent evidence has demonstrated that non-coding RNAs (ncRNAs) such as
small nucleolar RNAs (snoRNAs) and long non-coding RNAs (lncRNAs) can
act as master gene regulators at the transcriptional and
post-transcriptional epigenetic levels, and aberrant functions of these
RNAs are established hallmarks of tumorigenesis [[51]1, [52]2].
Notably, a number of RNA modifications, including ribosome RNA (rRNA)
2′-O-methylation (2′-O-Me), play an essential role in the regulation of
gene expression by altering and fine-tuning the properties of mRNAs,
rRNAs and lncRNAs [[53]3–[54]5]. Specifically, the most clearly
understood function of the C/D box small nucleolar RNAs (SNORDs) is
their ability to assemble small nucleolar ribonucleoprotein complexes
(snoRNPs) comprised of core RNP proteins (NOP58, NOP56, SNU13, FBL) and
rRNA such as SNORD27, which facilitates 2′-O-Me of A27 on 18S rRNA
[[55]6, [56]7]. The suppression of snoRNAs specific to U26, U44, and
U78 (corresponding to 28S-Am398, 18S-Am163, and 28S-Gm3745) reduced
rRNA modifications at the corresponding sites and led to severe
morphological defects and embryonic lethality, suggesting a critical
role for these rRNA 2′-O-Me events in vertebrate development [[57]8].
The endogenous mechanism responsible for regulating these snoRNAs and
their ability to mediate the modification-based regulatory network,
however, has not yet been thoroughly investigated in the context of
human tumorigenesis. Thus, it is crucial to clarify the expression
regulation pattern and the host gene lncRNAs and snoRNAs involved in
the progression and development of solid tumors.
Despite the discovery of snoRNAs in multiple species ranging from
bacteria to mammals in the 60’s, the functional role of these molecules
has remained enigmatic, and no cellular functions have been identified
until recently [[58]9, [59]10]. Recent studies have revealed the
significance and characteristics of snoRNAs in the context of
tumorigenesis. For example, overexpression of SNORD78 (C/D box) was
observed in non-small cell lung cancer and in hepatocellular carcinoma
[[60]11, [61]12]. Additionally, SNORD50A/B (C/D box) is deleted by
directly binding to Kras, and then affecting Kras expression across
multiple types of cancers [[62]13]. Increased SNORA42 (H/ACA box)
expression also serves as an independent prognostic factor for overall
survival times among cancer patients [[63]14]. SNORA55 (H/ACA box)
silencing in prostate cancer cell lines significantly inhibits cell
proliferation and migration [[64]15]. These findings highlight the
potential roles of snoRNAs in tumorigenesis, regardless of their C/D
box or H/ACA box classification. Despite the emerging knowledge
regarding the roles of snoRNAs in cancer, the expression landscape,
regulation network, and clinical relevance of snoRNAs have not been
systematically investigated in regard to cancer. These novel functions
require further clarification.
Of note, a number of the guide snoRNA-hosting genes in humans are
spliced, polyadenylated lncRNAs that are dynamically regulated during
cell proliferation, differentiation, and apoptosis [[65]16–[66]18].
These genes also exhibit cell- and tissue-specific expression patterns
[[67]19–[68]21]. snoRNA host genes are believed to be required for the
specific recruitment of snoRNPs that influence modification and
eventually re-enforce cell fate decisions to ensure a step-wise
developmental transition [[69]22]. For example, the host gene lncRNA
ZFAS1, which encodes three C/D box SNORD12 family members (SNORD12,
SNORD12B, SNORD12C), was observed to be significantly overexpressed in
a variety of human malignancies such as colorectal cancer, hepatic
cancer, and gastric cancer, etc. [[70]23–[71]27]. Based on this
information, it is likely that host gene lncRNAs play critical roles in
tumorigenesis and in clinical outcome. More importantly, most of the
snoRNAs appear to be the similar cellular localization with their host
genes and the snoRNPs complex, which suggests the possibility of
synergistically regulation function in tumorigenesis. The snoRNPs
assemble processes mainly include the ribosome biogenesis,
modification, and maturation, events that ultimately affect protein
translation fidelity [[72]28–[73]31]. Among these RNPs, NOP58 is an
adaptor of snoRNPs that binds to the conserved C box (RUGAUGA) and D
box (CUGA) of C/D box snoRNAs to provide a skeleton and bridge for the
entire snoRNP complex, ultimately maintaining the homeostasis of
epigenetic modifications [[74]5]. It must be noted, however, that
concrete examples of snoRNA and host gene co-regulation in response to
stimuli have not yet been reported. This prompted us to examine how
ribosome biogenesis and fine-tuning modification are controlled and to
determine whether and how this process is involved in the recruitment
of lncRNAs and snoRNPs.
In our current study, we demonstrate that the key motif of ZFAS1
directly interacts with the core component of snoRNPs/NOP58, promotes
NOP58 recruitment, and accelerates SNORD12C and SNORD78 snoRNPs
assembly to allow for the guiding of 2′-O-Me of 28S rRNA to specific
sites (Gm3878, Gm4593). Specifically, ZFAS1 knockdown results in
decreased RNA stabilization of NOP58, SNORD12C/78, and their 2′-O-Me
modification guidance, and this subsequently inhibits colorectal cancer
(CRC) cell proliferation and invasion and promotes cell apoptosis in
vitro and in vivo. Strikingly, SNORD12C or SNORD78 inhibition decreased
2′-O-Me modification regulated by ZFAS1, and this inhibited the RNA
stability and translation activity of their downstream targets such as
EIF4A3, LAMC2, and others. Based on this, we identified a previously
unrecognized signaling axis involving
ZFAS1-NOP58-SNORD12C/78-EIF4A3/LAMC2 that functions in CRC
tumorigenesis and our findings shed new light on our understanding of
lncRNAs-snoRNPs-mediated rRNA 2′-O-methylations in CRC tumorigenesis
and development.
Methods
Additional experimental details are included in Additional file [75]2.
Collection of the tissue specimen
In this study, human tissue samples were obtained from 157 patients
with colorectal cancer, who underwent surgical treatment at the
Department of General Surgery of the First Hospital of China Medical
University, Department of Medical Oncology of Cancer Hospital of China
Medical University between September 2014 and September 2015. This
study was approved by the Medical Ethics Committee of China Medical
University. All enrolled patients signed the written informed consent
form according to the relevant regulations. The tissues of CRC and
matched adjacent-tumor controls were snap frozen immediately in liquid
nitrogen after separated and stored at − 80 °C before using. The
inclusion, exclusion criteria, as well as clinicopathological data
collection and follow up of the included patients were described in the
Additional file [76]2.
LncRNAs-snoRNAs microarray assay
GeneChip® Human Transcriptome Array 2.0 (HTA2.0, Affymetrix, USA) was
selected and the microarray hybridization, data acquisition were
explored by Shanghai OE Biotech Technology Co, Ltd. (Shanghai, China).
The raw data have been deposited in Gene Expression Omnibus under an
accession number [77]GSE137511. This HAT2.0 designed array contains
more than 6.0 million distinct probes covering coding and non-coding
transcripts, and covered more than 285,000 full-length transcripts,
more than 245,000 coding transcripts, 40,000 non-coding transcripts and
339,000 probes covering exon-exon junctions of the human genomes. These
databases such as Ensembl, UCSC, NONCODE, RefSeq, lncRNAdb, Vertebrate
Genome Annotation (Vega), Mammalian Gene Collection (MGC), Human Body
Map lincRNAs, as well as related literatures were used to annotate the
determined transcripts. The data were analyzed with Robust Multichip
Analysis (RMA) algorithm using Affymetrix default analysis settings and
global scaling as normalization method, detail shown in Additional file
[78]2.
Gene expression analysis
Genesrping software (version 13.1; Agilent Technologies) was employed
to perform the raw data analysis. Deferentially expressed genes were
then identified through fold change as well as P value calculated with
t-test. The threshold of up- and down-regulated genes was set at fold
change ≥2.0 and P value ≤0.05. Afterwards, gene ontology (GO)
enrichment analysis and Kyoto Encyclopedia of Genes and Genomes (KEGG)
analysis were applied to determine the roles of these deferentially
expressed mRNAs played in these GO terms or pathways. Finally,
Hierarchical Clustering was performed to display the distinguishable
genes’ expression pattern among the included 6 samples.
Cell lines and cell culture
All of the human normal intestinal epithelial cell line HIEC and CRC
cells including HCT116, SW480, SW620, and HT29 were obtained from
Peking Union Medical College Cell Resource Center (PUMCCRC, Beijing,
China). The cells were cultured and maintained under standard cell
media and conditions. Specifically, HCT116 were maintained in RPMI-1640
(BI, Israel), SW480 and SW620 cells were grown in L15 (HyClone, USA),
HT29 were in McCoy’s 5A (BOSTER Biotech, China), and HIEC were cultured
in Dulbecco’s modified Eagle’s medium (DMEM, Invitrogen, USA) plus 10%
(v/v) fetal bovine serum (FBS), and 1% penicillin-streptomycin
(Invitrogen, USA). HEK293T cells (from PUMCCRC) were maintained in DMEM
plus 10% FBS. All of these cells were grown at 37 °C with a 5% CO[2]
cell culture incubator. In this study, all of the cells used were
genotyped by STR analysis and determined routinely for Mycoplasma
contamination.
Cell transfection
The plasmid extraction kit was purchased from Sangon Biotech (Shanghai,
China). All of the shRNA and overexpressing ZFAS1, NOP58 plasmids were
described in Additional file [79]2, and the plasmids nucleotide shRNA
sequences were listed in Table [80]S1 and Table [81]S2. Cells were
plated on 6-well plates to 60–70% confluence and transfected with
1μg/ml Lipofectamine 3000 (Invitrogen, Carlsbad, CA, USA) according to
the manufacturer’s instructions.
Reverse transcription-PCR (RT-PCR) assays and qPCR assays
According to the manufacturer’s instructions (see detail in Additional
file [82]2). Total RNA was extracted from tissues or cells by using
TRIzol reagent (Invitrogen, USA). For the RT-PCR assay, the reverse
transcription was performed from RNA to cDNA and PCR analyses were
performed by a PrimeScript™ RT-PCR Kit (Takara, Japan). Quantitative
real-time PCR (qPCR) was determined by SYBR Green I mix (Toyobo, Japan)
in triplicate based on an Applied Biosystems 7500HT Real-Time PCR
System. The mRNA relative expression was normalized to reference genes
GAPDH and/or U6. The reaction assays and primers used for qPCR were
listed in Table [83]S3 and Table [84]S4.
Cell proliferation assays
Transfected HCT116 and SW620 cells were seeded in 96-well plates
(100 μl/well) at the density of 5 × 10^3 cells/well for 24 h. Cell
viability was determined for 24, 48, 72 and 96 h by a Cell Counting
Kit-8 (CCK8, Bestbio, China) according to manufacturer’s instructions.
The absorbance of each well were measured and obtained the OD values at
490 nm with a microtiter plate reader (BioTek, USA). Each time point
was assayed in triplicate, and the experiment was replicated 3 times.
Flow cytometry assays
Cells were harvested and washed twice with cold 1 × PBS. For cell cycle
arrest analysis, cells were fixed with 70% ethanol and stored at 4 °C
overnight. After re-hydration with PBS, cells were treated with 20 μl
of RNase A (2 μg/ml), and incubated at 37 °C for 30 min. Cells were
then stained with propidium iodide (PI, 50 μg/ml) for 1 h at 4 °C. For
cell apoptosis analysis, cells were re-suspended with 100 μL of 1×
Annexin V binding buffer, and incubated with 5 μL of Annexin V-PE for
15 min and 5 μL of 7-AAD for 5 min in a darkroom at room temperature.
Finally, cells were analyzed by FACScalibur flow cytometer (BD, USA).
Western blot analysis
Cells were harvested and lysed by 1 × SDS buffer. Lysates were
sonicated and centrifuged (13,000 rpm, 4 °C) for 10 min. Proteins were
separated by 8–12% SDS-PAGE and transferred to polyvinylidene fluoride
membranes (Millipore, Bedford, MA). Membranes were immunoblotted with
anti-rabbit NOP58 (1:1000), EIF4A3 (1:1000) (Proteintech, Chicago, USA;
Abcam, UK) and anti-mouse LAMC2 (1:500), GAPDH (1:2000) (Abcam, UK;
Zsbio, Beijing, China), and then were incubated with hybrid secondary
antibody, and the data was collected by FluorChem V2.0 (Alpha Innotech
Corp, USA).
Immunofluorescence
Cells were grown on cover slides, fixed, and stained with indicated
antibodies. Antibodies used for immunofluorescence were as follows:
NOP58 (bs-19318R, 1:100, Bioss, Beijing, China), Alexa Fluor
anti-rabbit IgG (#4412, 1:500, Cell Signaling Technology). Cell nuclei
were counterstained with DAPI (Beyotime, Shanghai, China). Image
acquisition was performed on a confocal laser scanning microscope under
a 40 × objective (Nikon, Japan).
Co-localization of LncRNA/snoRNA and protein expression
Cells were cultured on cover slides and fixed normally following the
steps of immunofluorescence. Then RNA in situ hybridization was also
performed following the kit instructions above except counterstaining
with 0.1% Hematoxylin. Next, the cell was continued to stain with
indicated NOP58 antibody, Alexa Fluor anti-rabbit IgG and DAPI as
immunofluorescence. Similarly, Nikon C2 plus confocal microscope were
used to obtain images under a 40 × objective (Nikon, Japan).
Tissue microarray (TMA) and immunohistochemistry (IHC)
TMA and IHC method were performed as previously described with brief
modification [[85]32]. Briefly, the sections (4 μm) were deparaffinized
with xylene, rehydrated in a graded alcohol series, and washed in
distilled water. Then, sections were incubated in primary antibody of
NOP58 (bs-19318R, 1:100, Bioss, Beijing, China) overnight at 4 °C,
followed by incubation with biotinylated secondary antibodies for
30 min at 37 °C. The slides were incubated with horseradish peroxidase
coupled streptavidin for an additional 30 min (LSAB kit; Dako,
Glostrup, Denmark), and stained with DAB (3, 3-diaminobenzidine).
Sections were counterstained with hematoxylin, dehydrated, and mounted.
Protein expression levels were observed and counted under a microscope
(Eclipse 8i, Nikon, Japan), and the evaluation analysis was described
in Additional file [86]2.
RNA in situ hybridization (ISH) assay
In situ hybridization was performed strictly following the kit
instructions (Boster, Wuhan, China). Before prehybridized in
prehybridization solution at 42 °C for 2 h, slides were deparaffinized
and deproteinated, then incubated with a digoxin-labeled probe solution
(Dilute 4 times with 1 × PBS) at 37 °C over night (Specific probe
sequences were shown in Table [87]S5). After stringent washing, the
slides were exposed to a streptavidin- peroxidase reaction system and
stained with DAB (Zsbio, Beijing, China) for 2 min. Then 0.1%
Hematoxylin (Solarbio, Beijing, China) was used to counterstain the
slides for 5 min. ZFAS1 expression levels were observed and counted
under a microscope (Nikon, Tokyo, Japan), and the evaluation analysis
was described in Addition file 2.
RNA stability assay
SW620 cells were transfected with shZFAS1#1, ASO-SNORD12C followed with
a treatment by actinomycin D (ActD, CAS#:A4262, Sigma) at a final
concentration of 5 μg/mL for 0.5, 1, 2, 3, and 6 h. Total RNA was
extracted and analyzed by qRT-PCR. Then, the calculation of RNA
turnover rate and half-life (t[1/2]) of SNORD12C, SNORD78, NOP58,
EIF4A3, and LAMC2 were determined according to the previous
publications [[88]33]. Since ActD treatment results in transcription
stalling, the change of RNA concentration at a given time (dC/dt) is
proportional to the constant of RNA decay (K[decay]) and RNA
concentration (C) as shown in the following equation:
[MATH: dC/dt=−kdecayC :MATH]
Thus the RNA degradation rate k[decay] was estimated by:
[MATH: lnC/C0=−kdecayt :MATH]
When 50% of RNA is decayed (i.e., C/C[0] = 1/2), the equation below can
be used to calculate the RNA half-life (t[1/2]):
[MATH: ln1/2=−kdecayt1/2 :MATH]
From where:
[MATH:
t1/2=
ln2/kdecay :MATH]
Translation inhibition assay
Briefly, the SW620 cells were cultured for one dish at each time point
and then transfected with ASO-Sramble, ASO-SNORD12C and ASO-SNORD78.
After 48 h, the cells were treated with translation inhibitor,
cycloheximide (CHX, CAS#:C7698, Sigma) with a concentration of
200 μg/ml in the fresh cell medium. Thereafter, the cells were
incubated with CHX based on the different time points (0, 1, 2, 3, 4, 6
and 9 h). The zero hour represents the start time of treatment with
CHX. The total protein was isolated according to the time courses.
Finally, the expression levels of LAMC2, EIF4A3 were measured with
GAPDH as the internal control assayed by western blot method.
RNA pull-down assay
Briefly, biotin-labelled ZFAS1 oligonucleotide (probe sequence shown in
Table [89]S6) were conjugated to Streptavidin agarose resin beads.
Then, the ZFAS1-conjugated streptavidin beads were then incubated with
nuclear extract in binding buffer at 4 °C overnight. After washing with
1 × binding buffers, RNA-protein complexes were dissolved in 1× SDS
buffer, and analyzed by western blot assay (see Additional file [90]2
for details).
RTL-P assay for rRNA 2′-O-methylation
For the detection of 2′-O-methylation of SNORD12C and SNORD78, RT-PCR
was conducted referenced the previous publications with some
modifications [[91]34], RT was conducted in 25 μl reaction cocktails
with 100 ng of total RNA, 50 μM specific RT primers were denatured at
70 °C for 10 min, and then placed on ice, shown in Table [92]S7 and
Table [93]S8. Next, the RT buffer, 200 U M-MLV reverse transcriptase
(Takara), 40 U RNasin Ribonuclease inhibitor (Takara) and a low (10 μM)
or high (1 mM) concentration of dNTPs were mixed with an initial
annealing step at 42 °C for 1 h and then heated at 70 °C for 15 min.
Then the PCR reaction was determined and the PCR products were
separated on 2% agarose gels, and visualized by UV-trans-illumination.
Double-stranded primer based on single-stranded toehold (DPBST) assay
Total RNA was extracted from cells by using TRIzol reagent (Invitrogen,
USA). The condition of RNA reverse transcription to cDNA was modified
by adding low or high dNTPs in a Reverse Transcription Kit (Takara,
Japan). The specific primer was named as double-stranded primer based
on single-stranded toehold. The BST dsPrimers and reaction assays of
SNORD12C and SNORD78 were listed in Table [94]S9 and Table [95]S10.
Finally, qPCR assay was determined and the CT curves was obtained using
TB Green premix Ex TaqII (Takara, Japan) in triplicate.
Xenograft mice experiment
All protocols used followed the Regulations of Experimental Animal
Administration issued by the Ministry of Science and Technology of the
People’s Republic of China. The 4-week-old BALB/c-nu mice were
purchased from Shanghai Laboratory Animal Center (Shanghai, China).
Before the experiments, the mice were acclimatized to the new
environment for one week. 5 × 10^6 HCT116 cells were subcutaneously
injected into the right armpit region. When the tumors were visible,
the mice were randomly divided into four groups. The weight and tumor
size of the mice were measured every 5 days. Simultaneously, the
survival of mice was tracked and recorded. About 5 weeks after
injection, half of the mice in each group were sacrificed and the
subcutaneous tumors were isolated and measured, the rest was observed
for survival until the sixtieth day. Also, the tumor tissues were fixed
in 10% formalin for further research.
Statistical analysis
All of the statistical analysis was employed using SPSS 19.0 software
package (SPSS Inc. Chicago, USA), and GraphPad Prism 7.0 software
(GraphPad, USA). The data are presented as mean ± standard deviation
(s.d.) or median (quartile). Student’s t-test or Wilcoxon T-test was
performed to analyze the significant differences of the paired and
unpaired continuous variables. Pearson χ^2or Fisher’s exact test was
conducted to analyze the expression or distribution differences of the
variables. Kaplan-Meier method, Log-rank test, and univariate Cox
proportional hazard regression analysis were used to estimate the
potential prognosis associated indicators. P-values were two sides, and
P < 0.05 was considered statistically significant in all tests.
Results
Dysregulated lncRNAs, snoRNAs, and their co-expressing genes
To explore the dysregulated lncRNAs-snoRNAs-mRNAs and their
correlations and co-expression enrichment network, we performed
differential expression profiling analyses based on Affymetrix GeneChip
microarray that included three CRC patient tissues samples and their
matched tumor-adjacent normal tissues (n = 3). Significant differences
between these two groups were indicated by a ≥ 2-fold change and
P-value < 0.05, as illustrated in Fig. [96]1 a, b. The expression
differences for ncRNA (lncRNAs, snoRNAs) and mRNA were distributed
widely across all chromosomes, including the sex chromosomes (X and Y)
(Fig. [97]1a). In total, we identified 739 dysregulated ncRNAs,
including 654 lncRNAs and 85 snoRNAs, and 1164 dysregulated mRNA in
this cohort (Fig. [98]1a). Specifically, 293 up- and 361 down-regulated
lncRNAs, 62 up- and 23 down-regulated snoRNAs, and 469 up- and 695
down-regulated mRNAs were identified in this study that examined in CRC
tissues and matched tumor-adjacent tissues (Fig. [99]1b). The heatmap
clustering analysis of the top 30 dysregulated lncRNA expression
profiles revealed that ZFAS1 is dramatically upregulated (Log[2] Fold
Change/FC = 6.65), and the snoRNAs such as SNORD12C and SNORD78 was
remarkably up-regulated, where the Log[2] FC values were 5.71 and 6.86,
respectively (Fig. [100]1c, Table [101]S11-[102]S12). Additionally, a
dramatically higher expression level of ZFAS1 was observed in the vast
majority of cancers, particularly in CRC
([103]http://www.cbioportal.org/), based on the TCGA data (Fig.
[104]S1a). Furthermore, the Top 9 of up-regulated snoRNAs were selected
as candidate indicators including SNORD87, SNORD27, SNORD47, SNORD72,
SNORD75, SNORD78, SNORD12, SNORD12B, and SNORD12C based on our
microarray analysis (n = 3). Thereafter, the expression levels of those
indicators were identified in four CRC cell lines including HCT16,
SW620, SW480, HT29 and normal intestinal epithelial HIEC cells detected
by qPCR assay. Of interest, the expression levels of SNORD12C and
SNORD78 dramatically elevated compared with other candidate snoRNAs in
these included CRC cells, illustrated in Fig. [105]1d, and Fig.
[106]S1d.
Fig. 1.
[107]Fig. 1
[108]Open in a new tab
The correlation between ZFAS1, SNORD12C and SNORD78 expression in CRC
paired tissues and TCGA. a, A circular diagrams from the most inner
circle to the most outer circle represent the log2 fold change value of
down-regulated or up-regulated differentially expressed LncRNAs,
snoRNAs, and mRNAs of the CRC compared with matched adjacent-tumor
control tissues (n = 3, P < 0.05), the gene expression value of matched
adjacent-tumor control tissues, the gene expression value of CRC, the
different chromosome location of the genes in different colors, and the
ruler of chromosome. b, The volcano plot of LncRNAs, snoRNAs, and mRNAs
expression among included 3 pairs of CRC tissues and adjacent-tumor
control tissues (n = 3, Log 2 FC = 2.0, P < 0.05). c, Hierarchical
cluster heat map illustrating the most differentially expressed LncRNAs
and snoRNAs in CRC and corresponding paired adjacent-tumor control
tissues, selected top 30 up-regulated or down-regulated genes (n = 3,
P < 0.05). Red in heat map denotes upregulation. Blue denotes
downregulation. d, The expression levels of screened snoRNAs in normal
intestinal epithelial HIEC cell and CRC cells including HCT116, SW620,
SW480, and HT29 detected by qRT-PCR assays. Values are the mean ± s.d.
of n = 3 independent experiments. e, The Venn plot showing the
co-expression genes of ZFAS1, SNORD12C and SNORD78 in CRC microarray
enrichments (n = 3) in our CRC cohort. f, The schematic diagram of
2′-O-methylation of ribosomal RNA catalyzed by C/D box snoRNP
complexes. g, GO pathways enrichment analysis of the up-regulated
co-expression genes of ZFAS1, SNORD12C and SNORD78, and the related
biological functions. h, The linear correlation analysis representing
the relation of NOP58 expression levels with ZFAS1 upon TCGA CRC
database. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001
To further investigate the potential functions of ZFAS1 expression and
its related snoRNAs in CRC patients, we conducted the intersection of
the potential target mRNAs, which were enriched from the up-regulated
co-expressed with ZFAS1 and up-regulated co-expressed with SNORD12C and
SNORD78 by the Bioinformatics & Evolutionary Genomics platform
([109]http://bioinformatics.psb.ugent.be/webtools/Venn/). Subsequently,
202 potential target genes were intersected among these interaction
networks, and these networks mainly consisted of components of the C/D
box small nucleolar ribonucleoproteins (snoRNPs) such as NOP58 (Fig.
[110]1e, f, Table [111]S14). Based on the TCGA dataset (n = 638), NOP58
expression levels were also significantly increased in the CRC tissue
samples compared with those in the healthy donors (n = 51), which was
in contrast to levels of other snoRNPs such as NOP56, SNU13, and FBL
(Fig. [112]S1b). Importantly, heat map clusters revealed that NOP58 was
dramatically up-regulated in the included CRC tissues and the matched
tumor-adjacent normal tissues (Fig. [113]S1c, Table [114]S13). As
expected, the Top 2 results of the Gene Ontology (GO) analysis
indicated that the potential functions were focused primarily on the
regulation of ribonucleoprotein complex biogenesis and
ribonucleoprotein complex assembly (Fig. [115]1g). Further correlation
analyses verified that NOP58 showed stronger associated with the
expression of ZFAS1 than those indicators such as NOP56, SNU13, and FBL
(Fig. [116]1h, Fig. [117]S1e, Table [118]S14).
Correlations among the levels of ZFAS1, SNORD12C, SNORD78, and NOP58 in CRC
cells
To further demonstrate the potential influence of ZFAS1 expression on
SNORD12C, SNORD78, and NOP58, we detected the expression levels of
these genes in 4 CRC cells (HT29, HCT116, SW480, and SW620) and in
normal control HIEC cells. Our results revealed that ZFAS1 was
expressed at a high level, and this expression was accompanied by
elevated expression of SNORD12C, SNORD78, and NOP58 in CRC cells as
indicated by both RT-PCR and qPCR assays (Fig. [119]2a, b). Similar
results were obtained from 30 pairs of CRC tissues and matched
tumor-adjacent normal tissues, where an elevated expression of ZFAS1 in
CRC tissues and a further consistently up-regulated of SNORD12C,
SNORD78, and NOP58 were detected by RT-PCR and qPCR assays (Fig.
[120]2c, Fig. [121]S2). Importantly, the linear regression analysis
further identified the positive correlation among those indicators in
our included CRC tissues and controls, shown in Fig. [122]S3. These
findings suggested that ZFAS1 expression was positive correlated with
the expression of SNORD12C, SNORD78, and NOP58 in CRC cells and
tissues. More importantly, co-localization assays using ISH combined
with IF revealed that the majority of ZFAS1, SNORD12C, and SNORD78 were
co-distributed and co-located with NOP58 within the cell nucleus of the
HCT116 cells (Fig. [123]2d). Additionally, the ectopic ZFAS1 expression
caused a substantial elevation in NOP58 and SNORD12C/78 expression
levels in HCT116 and SW620 CRC cells (Fig. [124]2e). In contrast, ZFAS1
knockdown significantly inhibited the expression levels of NOP58 and
SNORD12C/78 in CRC cells (Fig. [125]2f).
Fig. 2.
[126]Fig. 2
[127]Open in a new tab
The expression of ZFAS1, SNORD12C, and SNORD78 and subcellular
localization. a and b, Expression levels of ZFAS1, NOP58 (a) and
SNORD12C or SNORD78 (b) in CRC cell lines including HT29, HCT116,
SW480, and SW620 and normal intestinal epithelial cell line HIEC
detected by RT-PCR and qPCR assay. Values are the mean ± s.d. of n = 3
independent experiments. c, Representative images of ZFAS1, NOP58,
SNORD12C and SNORD78 expression in paired CRC and matched adjacent
normal tissues determined by RT-PCR assays (5 representative data was
shown). d, Co-localization of ZFAS1, SNORD12C and SNORD78 with NOP58
protein detected by ISH and IF assays in HCT116 cells. Scale
bar = 5 μm. e and f, The expression of ZFAS1, NOP58, SNORD12C, and
SNORD78 after overexpression or silencing ZFAS1 in both of HCT116 and
SW620 cells by qRT-PCR assay.*, P < 0.05; **, P < 0.01; ***, P < 0.001;
****, P < 0.0001
Taken together, these data indicated that ZFAS1 expression was
up-regulated companied by its correlated SNORD12C/78 and NOP58 in human
CRC cells and tissues. A positive regulation pattern was further
identified between ZFAS1 and SNORD12C/78 and NOP58 in paired CRC
tissues and by interfering ZFAS1 expression in CRC cells.
Effect of ZFAS1 on RNA stability and rRNA 2′-O-methylation
Based on the observation that the knockdown of ZFAS1 inhibits the
expression of snoRNAs, SNORD12C/78, and NOP58, we next investigated if
the changes in the expression levels of mRNAs and snoRNAs after ZFAS1
silencing were due to accelerated mRNA decay. To achieve this, we
conducted the RNA stability assays in SW620 cells with the treatment of
ActD. As expected, the half-lives of RNAs remaining significantly
decreased in those indicators including NOP58, SNORD12C, and SNORD78
compared with the NC group (Fig. [128]3a). Furthermore, two methods
including RTL-P and DPBST assays were explored to confirm the 28S rRNAs
2′-O-Me activity mediated by SNORD12C or SNORD78. Notably, RTL-P assays
demonstrated that ZFAS1 knockdown dramatically reduced the 28S rRNA
2′-O-Me activity at the G3878 site modified by SNORD12C and the G4593
site modified by SNORD78 under the lower dNTPs concentrations, however,
no significant difference was observed under the higher dNTPs
conditions (Fig. [129]3b, Fig. [130]S3f, g). Similarly, DPBST assays
further revealed that the 2′-O-Me activities modified by SNORD12C or
SNORD78 were significantly decreased upon ZFAS1 knockdown, meanwhile,
overexpression of ZFAS1 elevated the activities of 2′-O-Me (Fig.
[131]3c-f).
Fig. 3.
[132]Fig. 3
[133]Open in a new tab
ZFAS1 inhibits RNA stability of SNORD12C, SNROD78, and the specific
sites of rRNA 2′-O-methylation. a, Reducing NOP58, SNORD12C, and
SNROD78 half-life (t[1/2]) by silencing ZFAS1 in SW620 cells. Values
are the mean ± s.d. of n = 3 independent experiments. b, The rRNAs
2′-O-Me activities of G3878 site and G4593 site were decreased after
ZFAS1 knockdown in SW620 and HCT116 detected by RTL-P assay. c, The
schematic structures showing a novel method called double-stranded
primer based on single-stranded toehold (DPBST) assay for detecting
rRNAs 2′-O-Me levels. d and e, The 28S rRNA G3878 and G4593 sites of
2′-O-Me mediated by SNORD12C or SNORD78 at the high or low dNTPs
conditions after silencing ZFAS1 (Upper) or overexpression ZFAS1
(Lower) in SW620 cells detected by DPBST assays. f, The DPBST detecting
statistical results of 2′-O-Me by qPCR assay. *, P < 0.05; **,
P < 0.01; ***, P < 0.001; ****, P < 0.0001
Taken together, our results indicate that ZFAS1 exerts a significant
effect on the RNA stability and 2′-O-Me activities mediated by SNORD12C
or SNORD78, ultimately affecting CRC tumorigenesis and epigenetic
2′-O-Me levels of target 28S rRNAs.
Effects of ZFAS1 on CRC cell proliferation and apoptosis
To further verify the impact of ZFAS1 on the biological mechanisms of
CRC, we used CCK8 assays to determine that overexpression of ZFAS1
significantly enhanced the proliferation of HCT116 and SW620 cells,
while knockdown of ZFAS1 suppressed the proliferative ability of these
two cell lines (Fig. [134]4a). Additionally, knockdown of ZFAS1 greatly
decreased the migrated cells numbers in both HCT116 and SW620 cells,
and in contrast, the ectopic expression of ZFAS1 promoted an increase
in the numbers of HCT116 and SW620 cells (Fig. [135]4b). ZFAS1
knockdown also resulted in a marked increase in cell apoptosis, and
overexpression of ZFAS1 substantially inhibited apoptosis in both
HCT116 and SW620 cells as determined by flow cytometry (Fig. [136]4c,
Fig. [137]S4a). RT-qPCR analyses indicated that the inhibition of ZFAS1
expression positively affected NOP58 expression in both HCT116 and
SW620 cells, and similarly, NOP58 protein expression levels were also
decreased after ZFAS1 inhibition in HCT116 and SW620 CRC cells as
determined by WB and IF assays (Fig. [138]4d, e, Fig. [139]S4 b, c).
Similarly, in vitro rescue experiments revealed that NOP58
overexpression reversed the ZFAS1 inhibition effect on CRC molecular
characteristics including cell proliferation ability, cell apoptotic
rates in both HCT116 and SW620 cells assayed by CCK8 (Fig. [140]S4d),
and flow cytometry analysis (Fig. [141]S4e). We next used tissue
microarray (TMA) and immunohistochemistry (IHC) to examine a relatively
large number of samples that included 157 pairs of CRC and matched
tumor-adjacent normal tissues (Fig. [142]4f). The cut-off value of
ZFAS1 or NOP58 expression was determined based on the ROC curve method
(Fig. [143]S5 a, b). Consistent with the results in CRC cells, the
expression levels of ZFAS1 and NOP58 were elevated in the CRC tissues
compared to the levels detected in the tumor-adjacent normal tissues
(Fig. [144]4g), and a large positive linear correlation pattern was
confirmed to exist between ZFAS1 and NOP58 within this cohort (Fig.
[145]S5c). As expected, the prognostic analysis revealed that elevated
ZFAS1 expression significantly correlated with shortened overall
survival (OS, P = 0.002) and reduced disease-free survival (DFS,
P = 0.002) (Fig. [146]4h, Fig. [147]S5d). Consistently, higher
expression of NOP58 was significantly associated with poor OS
(P = 0.006) and DFS (P = 0.006) (Fig. [148]4h, Fig. [149]S5d). Further
multivariate Cox regression analyses also confirmed the prognostic
values of ZFAS1 and NOP58 after adjusting for confounders including age
and pathological pattern at diagnosis for OS and tumor stage for DFS
(Table [150]S15-[151]S16).
Fig. 4.
[152]Fig. 4
[153]Open in a new tab
Effects of ZFAS1 on cell proliferation and apoptosis by regulating
NOP58 expression in CRC cells. a, CCK8 assays were used to identify the
cell proliferation abilities upon ZFAS1 silencing or overexpressing in
HCT116 and SW620 cells. n = 3 independent experiments. b, The migrated
cell numbers were determined after ectopic or knockdown ZFAS1 in HCT116
and SW620 cells. c, The percentage (%) of cell apoptosis was detected
upon ZFAS1 overexpressing or silencing in HCT116 and SW620 cells by
Flow cytometry. n = 3 independent experiments. d and e, The NOP58
expression was conducted after overexpressing or knocking down ZFAS1 by
Western blot (d), and IF assays (e). f, ISH and IHC methods were used
to determine the cell localization and expression levels of ZFAS1 and
NOP58 in CRC tissues and matched adjacent-tumor controls (n = 157).
Scale bars = 100 μm. g, Violin charts displaying the expression scores
and levels of ZFAS1 and NOP58 between CRC tissues and matched paired
adjacent normal tissues (n = 157). Nonparametric tests and Median and
95%CI were shown. h, Kaplan-Meier curves representing the impact of
ZFAS1 and NOP58 on overall survival in this included CRC cohort.*,
P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001
Collectively, our results indicated that overexpression of ZFAS1
significantly accelerated the progression of human CRC cells by
promoting the expression of NOP58 levels; however, the molecular
mechanisms and underlying functions require further investigation.
NOP58 recruitment was accelerated upon ZFAS1-mediated 2′-O-me activities of
SNORD12C/78
To further clarify the molecular mechanism by which ZFAS1 affects the
2′-O-Me activities modified by SNORD12C/78 and its regulation pattern,
the catRAPID platform ([154]http://service.tartaglialab.com) was
employed to evaluate the interaction propensity and discriminative
power between the ZFAS1 nucleotide index and the NOP58 residue index
(Fig. [155]5b). As expected, ZFAS1 exhibited significant interaction
propensity (value = 209) and discriminative power (100%) for NOP58. The
critical binding motif were predicted by POSTAR2
([156]http://lulab.life.tsinghua.edu.cn/postar/) (Fig. [157]5a).
Furthermore, MOE software was used to identify the direct binding
domain and the critical amino acids within NOP58 3D structure,
illustrated in Fig. [158]5c.Thereafter, the biotin-labeled LncRNA-ZFAS1
probes were synthesized containing the binding motif with NOP58
(ZFAS1-WT) or a corresponding mutant sequence (ZFAS1-Mut). RNA
pull-down assays further demonstrated that the ZFAS1-WT probe, but not
ZFAS1-Mut, significantly pulled down endogenous nuclear NOP58 protein
(Fig. [159]5d), and this enrichment was significantly reduced after
silencing ZFAS1 expression, indicating a direct regulation of ZFAS1
with NOP58 by specific binding manner. Next, NOP58 knockdown or
overexpression cell models were established by SW620 and HCT116 cells
(Fig. [160]5e). qPCR assays confirmed that NOP58 mRNA expression can be
recovered compared with the NC group after co-transfection with
pcDH-ZFAS1 and shNOP58 or shZFAS1#1/#2 and pcDH-NOP58 (Fig. [161]5f).
WB assays further determined that the decreased NOP58 protein
expression was retrieved by co-transfection with pcDH-ZFAS1 and shNOP58
or shZFAS1#1/#2 and pcDH-NOP58 in both SW620 and HCT116 cells (Fig.
[162]5g). More importantly, the 2′-O-Me activities mediated by SNORD12C
and SNORD78 were dramatically decreased after knockdown NOP58
expression in both SW620 and HCT116 cells (Fig. [163]5h, Fig. [164]S6a,
b). Similar results from the rescue experiments demonstrated that the
2′-O-Me activities were recovered both in HCT116 and SW620 cells as a
result of NOP58 overexpression after co-transfected with shZFAS1#1/#2
(Fig. [165]5i, Fig. [166]S6c, d).
Fig. 5.
[167]Fig. 5
[168]Open in a new tab
ZFAS1 promotes rRNA G3878 and G4593 2′-O-methylation by targeting NOP58
protein. a, CLIP database showing the motif of ZFAS1 targeting NOP58
predicted by online software
([169]http://lulab.life.tsinghua.edu.cn/postar/index.php). b,
Bioinformatics online software predicting the specific binding sequence
and sites of ZFAS1 secondary structure and NOP58 protein
([170]http://www.tartaglialab.com/). c, MOE multi-functional docking
platform showing the specific docking sites between ZFAS1 tertiary
structure and NOP58 protein. d, RNA pull-down followed by western blot
showed in vitro binding of the ZFAS1-Wild, ZFAS1-Mutant, and antisense
RNA probes with NOP58 protein after ZFAS1 silencing in SW620 cells. The
biotin labeled probes are presented in Table [171]S6. e and f, The
NOP58 expression after overexpressing or knockdown NOP58 by qRT-PCR
assay (e), and ZFAS1 rescued NOP58 mRNA expression after co-transfected
ZFAS1 and NOP58 (f). g, Western blot assays detected the NOP58 protein
expression after overexpressing, knocking down NOP58, as well as
co-transfected ZFAS1 and NOP58 vectors. h and i, The 28S rRNA G3878 and
G4593 2′-O-Me activities after overexpressing or knockdown NOP58 in
SW620 and HCT116 cells by RTL-P (h), and ZFAS1 rescued the 2′-O-Me
activities after co-transfected ZFAS1 and NOP58 (i).*, P < 0.05; **,
P < 0.01; ***, P < 0.001; ****, P < 0.0001
Taken together, these data suggest that ZFAS1 recruits NOP58 by direct
binding that is mediated by 2′-O-methylation activities of SNORD12C and
SNORD78, and this recruitment significantly impacts CRC tumorigenesis
and development.
SNORD12C/78-mediated 2′-O-me regulates the translation activity of target
genes in a ZFAS1- dependent manner
To further clarify the function of snoRNAs in the context of
ZFAS1-mediated regulation of CRC cell proliferation and the translation
of target genes, we employed enrichment of co-expression analyses
between SNORD12C/78 and mRNA expression to search the possible
downstream target genes, including EIF4A3, LAMC2, MACC1, and CSE1L,
responsible for regulating 2′-O-Me activities. We found that the
expression levels of EIF4A3 and LAMC2 expression were significantly
decreased upon knockdown of SNORD12C/78 expression as assessed by qPCR
and WB assays (Fig. [172]6a, b). Additionally, RNA stability assays
further identified that the half-life of LAMC2 and EIF4A3 mRNA was
reduced by knockdown SNORD12C expression or by treatment with ActD in
SW620 cells compared to the half-life in cells treated with empty
vector (negative control) (Fig. [173]6c). In agreement with these
findings, the protein expression levels of LAMC2 and EIF4A3 were
measured after treatment with the CHX, and we observed that the
expression levels gradually decreased throughout this time course (Fig.
[174]6d) in SW620 cells. Finally, the 2′-O-Me activities modified by
SNORD12C or SNORD78 were significantly decreased after silencing
SNORD12C or SNORD78 expression in both of SW620 and HCT116 cells (Fig.
[175]6e, Fig. [176]S7a, b). Similarly, this decrease was rescued by
ZFAS1 overexpression when co-transfected with ASO-SNORD12C or
ASO-SNORD78 assessed by RTL-P assays in both HCT116 and SW620 cells
(Fig. [177]6f, g), suggesting the existence of a 2′-O-Me mediated
regulation between ZFAS1 and SNORD12C or SNORD78.
Fig. 6.
[178]Fig. 6
[179]Open in a new tab
SNORD12C/78-mediated 2′-O-Me modification regulates target genes
expression in a ZFAS1-dependent manner. a and b, EIF4A3 and LAMC2 mRNA
expression was determined after knocking down SNORD12C or SNORD78 by
qRT-PCR assays(a), and Western blot method(b). c, Reducing EIF4A3 and
LAMC2 half-life by silencing SNORD12C treated by the ActD in SW620
cells. d, Reducing the translation activity of EIF4A3 and LAMC2 by
silencing SNORD12C and SNORD78 treated by the CHX in SW620 cells. e,
The 28S rRNA G3878 and G4593 2′-O-Me activities were declined after
silencing SNORD12C or SNORD78 expression in SW620 and HCT116 cells
detected by RTL-P assays. f, Overexpressing ZFAS1 rescued the 2′-O-Me
activities under the lower dNTPs conditions after silencing SNORD12C
and SNORD78. g, The statistical plot of overexpressing ZFAS1 rescued
the 2′-O-Me activities after silencing SNORD12C and SNORD78 *,
P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001
ZFAS1 inhibits cell proliferation by interacting with NOP58 in vivo
To evaluate the molecular mechanism of ZFAS1 interaction with NOP58 in
the context of CRC in vivo, we first established xenografts in BALB/c
nude mice by inoculating HCT116 cells that were stably co-transfected
with ZFAS1 and NOP58-WT, NOP58-Mut vector, or NC controls within the
same mice at their right armpits (Fig. [180]7a). The mice were
sacrificed, and the xenograft tumors were removed at the fifth week
after implantation or followed up until death. As expected, a dramatic
increase in the xenograft tumor weight and tumor volume was observed
after the co-transfection with ZFAS1 and the NOP58-WT vector compared
to that observed after transfection with NC or the overexpressing ZFAS1
vector (Fig. [181]7b, c, d), indicating that the promotion of
tumorigenic potential in CRC cells is a result of ZFAS1 specifically
interacting with NOP58 in xenograft mouse models. To further confirm
this, we compared the survival times among different groups, and we
observed a substantially reduced survival time in xenograft mice within
the ZFAS1 and NOP58-WT co-transfection group (Fig. [182]7e), indicating
the interaction between ZFAS1 and NOP58 acts as a synergistic risk
factor in the prognosis evaluation of CRCs. Additionally, in agreement
with in vitro data, SNORD12C/78, EIF4A3, and LAMC2 expression were
significantly up-regulated by the presence of ZFAS1 and NOP58 in vivo
as assessed by qPCR (Fig. [183]7f), suggesting a direct binding
regulated by ZFAS1. IHC and WB analyses further indicated that NOP58
was significantly increased in the group co-transfected with ZFAS1 and
the NOP58-WT vector, and this was accompanied by an increase in the
levels of their targets such as EIF4A3 and LAMC2 (Fig. [184]7g-i).
Collectively, these results demonstrated that ZFAS1 promotes the
proliferation and survival of CRCs by targeting NOP58 to modulate its
protein translation.
Fig. 7.
[185]Fig. 7
[186]Open in a new tab
ZFAS1 inhibited proliferation by targeting NOP58 protein in vivo. a,
Schematic diagram of xenografts in BALB/c nude mice by inoculating
HCT116 cells that were stably co-transfected with ZFAS1,
ZFAS1-NOP58-Wild, and ZFAS1-NOP58-Mut, as well as the control with
empty vector at their right armpits. Then half of the xenografts were
sacrificed at the 35th day after injection and the other half were
tracked until death. b, Mean tumor weight of each group xenografts in
nude mice. Data are shown as mean ± s.d., n = 6 for each group. c, Mean
tumor volumes on different days for each group xenografts in nude mice.
Data are showed as mean ± s.d., n = 6 for each group. d, Representative
tumors size excised on day 35 are shown. e, Kaplan-Meier graph showing
overall survival of each group, n = 6. f, qPCR assays were performed to
determine the (m) RNA expression of ZFAS1, NOP58, LAMC2, EIF4A3,
SNORD12C, SNORD78 in above each group. g, h and i, IHC assay and
western blot to detect the protein expression of NOP58 in xenografts
tumor tissues of each group. The groups were as follows: NC (empty
vector); ZFAS1 (pcDH-ZFAS1); ZFAS1 + NOP58-Wild (co-transfected with
pcDH-ZFAS1 and pcDH-NOP58-Wild); ZFAS1 + NOP58-Mut (co-transfected with
pcDH-ZFAS1 and pcDH-NOP58-Mut).*, P < 0.05; **, P < 0.01; ***,
P < 0.001; ****, P < 0.0001
Discussion
Colorectal cancer (CRC) remains one of the most common digestive system
malignancies that exhibit higher morbidity and mortality worldwide
[[187]35]. In humans, a complicated set of factors are involved in CRC
tumorigenesis, including environmental exposures, inherited genetic
mutations (particularly multiple epigenetic modifications), and other
factors [[188]36–[189]38]. Despite the use of advanced treatments such
as surgical resection, chemotherapy, and/or radiotherapy, the mortality
rate of CRC patients has not appreciably improved, and this is likely
due to deficiencies in effectively screening for molecular biomarkers.
Accumulating evidence indicates that the non-coding RNAs (ncRNAs),
specifically long non-coding RNAs (lncRNAs) and small nucleolar RNAs
(snoRNAs) of short ncRNAs, can act as master regulators of gene
regulation and RNA modification [[190]39, [191]40]. Defective function
of these molecules has become one of the hallmarks of tumorigenesis and
cancer development. Thus, insight regarding reliable molecular
biomarkers that are involved in the progression and development of CRC
is becoming more valuable, as it could considerably facilitate the
clinical diagnosis and promote the prognosis of CRC patients.
Ribose 2′-O-Me, the most prevalent internal chemical modification
between ribosomal RNA and snoRNAs, plays critical roles in many
bioprocesses such as RNA stability and translation fidelity in multiple
species ranging from bacteria to mammals [[192]41–[193]43]. Notably,
dysregulated levels of C/D box snoRNA correlated with alterations in
2′-O-Me of rRNA, and these alterations can contribute to cancer
progression and outcome [[194]44]. The underlying molecular mechanism
of the snoRNAs and their related regulation network remain elusive,
however, especially in colorectal cancer. Here, our studies identify
ZFAS1 and its direct interaction target NOP58 (the core components of
snoRNPs) as central to the master C/D box snoRNAs mediated
2′-O-methylation epigenetics modification network that is essential for
CRC initiation and maintenance (see the proposed model in Fig. [195]8).
Briefly, in normal cells, ZFAS1 and NOP58 (a direct binding target) are
normally expressed, and the C/D box snoRNAs SNORD12C and SNORD78
(specific assembly snoRNP complex) are recruited at normal levels. In
CRC cells and tissues, ZFAS1 and its endogenous recruiting protein
NOP58 are up-regulated, and this promotes the SNORD12C and
SNORD78-mediated 28S rRNAs 2′-O-Me activities at specific sites to
substantially increase the translation activation of downstream genes
such as EIF4A3, LAMC2, and others. This promotion can be blocked by
transfection with a NOP58-mutant vector in vivo, and this blockage
reverses the expression patterns of the above genes (Fig. [196]8).
Fig. 8.
[197]Fig. 8
[198]Open in a new tab
The schematic diagram illustrated that ZFAS1 targeting recruitment
NOP58 regulated rRNA 2′-O-methylation mediated protein translation to
mediate colorectal cell proliferation. In normal intestinal epithelial
cells, ZFAS1, NOP58 and snoRNAs are expressed normally, as well as the
level of the rRNA 2′-O-methylation and translation. In CRCs cells and
tissues, ZFAS1 is overexpressed, and its binding protein NOP58, which
is one of the snoRNPs, is also overexpressed. ZFAS1 increases the
snoRNPs formed by the synergistic recruitment of snoRNAs in combination
with NOP58, especially SNORD12C and SNORD78.This results in an increase
in the level of rRNAs 2′-O-Me modification, which ultimately leads to
changes in translational activity and precision of downstream target
genes, thereby mediating colorectal cancer proliferation
Increasing studies have provided evidence that lncRNAs (e.g., H19,
MEG3, MALAT1, etc.) exert their functions in the context of cancer
initiation and progression by influencing epigenetic modifications such
as DNA CpG methylation and RNA N6-methyladenosine (m6A); however, they
have not been observed to influence rRNA 2′-O-methylation
[[199]45–[200]47]. In the current study, we demonstrated that ZFAS1
accelerates the recruitment of SNORD12C and SNORD78 through a direct
interaction with NOP58, a core component of RNPs, to assemble the
corresponding snoRNP complexes. This subsequently promotes the 2′-O-Me
of 28S rRNA and ultimately plays an important role in CRC tumorigenesis
and clinical outcomes. Recently, critical lncRNAs were found to be
dysregulated and to act as oncogenes that influence cell fate decisions
by promoting cell proliferation, migration, invasion, and apoptosis
inhibition. Similar to the recent studies demonstrating the oncogenic
role of ZFAS1 in NSCLC, HCC, gastric cancer, and other cancers
[[201]48, [202]49]. For example, Zhang et al. reported that ZFAS1
involved in gastric cancer initiation and development based on ceRNA
network and Wnt/β-catenin signaling axis, which provide new targets and
biomarkers for gastric cancer treatment and evaluation [[203]50,
[204]51]. Consistently, our findings demonstrated that ZFAS1 functions
as a critical oncogene in CRC and that its expression/function is
required for both development and maintenance in CRC cells and tissues.
In contrast to the general assumption that noncoding SNORD-host
transcripts function only as vehicles to generate snoRNAs, ectopic
expression of ZFAS1 in CRC cells resulted in elevated cell
proliferation, invasion promotion, and cell apoptosis inhibition
[[205]22]. Additionally, ectopic expression of ZFAS1 also up-regulated
the levels of the SNORD12C (the host gene ZFAS1) and SNORD78, and
resulted in elevated NOP58 expression. Interestingly, knockdown of
ZFAS1 further resulted in decreased RNA stabilization of NOP58,
SNORD12C, and SNORD78 and in reduced levels of 28S rRNA 2′-O-Me at
specific sites (SNORD12C: 28S-Gm3878, SNORD78: 28S-Gm4593). Complete
distribution profiles of the residues revealed that the biological
characteristics of snoRNAs primarily depend upon the types that are
associated with a set of core RNPs to assemble stable and functional
snoRNP particles to thereby mediate the consequent biological
functions. C/D snoRNAs are associated with evolutionarily stable and
highly conserved core RNPs, including NOP58, NOP56, SNU13, and with the
methyltransferase FBL to guide 2′-O-Me of target rRNAs. Among these
proteins, NOP58, as an adaptor in snoRNPs, binds to the conserved C box
(RUGAUGA) and D box (CUGA) of C/D box snoRNAs to provide a skeleton and
bridge for the entire snoRNP complex, ultimately maintaining the
homeostasis of epigenetic modifications [[206]52]. Our current studies
further supported that NOP58 can directly interact with ZFAS1 to induce
its function, and this complex then participates in C/D box SNORD12C
and SNORD78 snoRNP complex assembly that is involved in 2′-O-Me of
nucleotides at specific positions in rRNAs that ultimately contribute
to CRC tumorigenesis. It is noteworthy that this pattern was confirmed
by using the ZFAS1-Mut probe and/or vector with the binding sequence,
indicating a direct binding interaction between ZFAS1 and NOP58 that
influenced regulation in vitro and in vivo.
2′-O-Me is present within various cellular RNAs and is essential for
RNA biogenesis and functionality [[207]34]. Although the importance of
rRNA 2′-O-Me was established by suppression of snoRNA expression in a
zebrafish model [[208]8], it has not been extensively studied in human
cellular translation. Here, we developed a novel method referred to as
DPBST (Double-stranded primer based on single-stranded toehold) to
detect 2′-O-Me activities in rRNAs. This method allows for precise
mapping and superior sensitivity compared to that of previous classical
methods such as RTL-P. Using RTL-P and DPBST assays, we further
investigated the molecular mechanisms of ZFAS1 in regard to the
biological functions of 2′-O-Me, including improvement in RNA
stability, fine modulation of its conformation, and its importance in
ribosomal translation. Indeed, the 2′-O-Me activities were
significantly decreased after treatment with ASO-SNORD12C/78 in CRC
cells. In contrast, this decrease was almost completely rescued by
ZFAS1 overexpression in cells co-transfected with ASO-SNORD12C/78.
Additionally, RNA stability and translation activity of their
downstream targets such as EIF4A3 and LAMC2 were significantly affected
in this regulatory network. Specifically, knockdown of SNORD12C/78
remarkably decreased the expression, the mRNA remaining half-life, and
the translation fidelity of EIF4A3 and LAMC2 in CRC cells. These data
indicated that ZFAS1 functioned as an endogenous regulator to increase
the expression of SNORD12C/78 by directly binding to NOP58. SNORD12C
and SNORD78 also correlated with altered 2′-O-Me activities within 28S
rRNA at corresponding sites (28S-Gm3878, 28S-Gm4593) that contribute to
cell fate by influencing downstream target protein synthesis and
translation fidelity in proteins such as EIF4A3 and LAMC2. We also
observed that these regulation patterns can be regulated by ZFAS1.
Based on these observations, our data provide new insights into the
involvement of lncRNA in the C/D box snoRNAs-mediated 2′-O-Me
modification field, ultimately providing a solid basis for future
development and for further understanding of pathological properties of
post-transcriptional RNA alterations, particularly in cancers.
In summary, our research provides insights into a novel molecular
mechanism of the lncRNA ZFAS1 in the regulation of CRC initiation and
pathogenesis. By direct binding to the core RNP NOP58, ZFAS1 promoted
SNOR12C/78 snoRNPs complex assembly and 28S rRNA 2′-O-methylation at
corresponding sites to substantially influence the RNA stability and
translation activity their downstream target genes (e.g., EIF4A3 and
LAMC2). This, in turn, mediates CRC cell proliferation promotion and
apoptosis inhibition in vitro and in vivo. Our study identified a
previously unrecognized signaling axis involving
ZFAS1-NOP58-SNORD12C/78-EIF4A3/LAMC2 that functions in CRC development
and progression. Therefore, our work sheds new light on the potential
applications of lncRNA, snoRNAs, and cellular
2′-O-methylation-dependent translation networks in the prevention and
therapy of CRCs.
Supplementary information
[209]12943_2020_1201_MOESM1_ESM.pdf^ (723.7KB, pdf)
Additional file 1: Table S1.Short hairpin RNAs (shRNAs) sequence
against ZFAS1. Table S2.Short hairpin RNAs (shRNAs) sequence against
NOP58. Table S3. The reaction assays of qRT-PCR method. Table S4.
Primers used in qRT-PCR assays. Table S5. Probes used in situ
hybridization (ISH) assay. Table S6. RNA probes used for RNA pull-down
assays. Table S7. Primers used in site-specific 2′-O-methylation for
RTL-P assay. Table S8. RTL-P assays for rRNA 2′-O-methylation. Table
S9. Probes used in site-specific 2′-O-methylation for double-stranded
primer based on single-stranded toehold (DPBST) assays. Table S10.
Double-stranded primer based on single-stranded toehold (DPBST) assays.
Table S11. Data of LncRNAs cluster in Heat map analysis. Table S12.
Data of snoRNAs cluster in Heat map analysis. Table S13. Data of mRNAs
cluster in Heat map analysis. Table S14. The enriched target genes
intersected by ZFAS1, SNORD12C/78 in GO analysis. Table S15. Prognostic
information of included colorectal cancer patients (n = 157). Table
S16. Multivariate COX regression analysis of the association of ZFAS1
and NOP58 expression with DFS and OS in CRC patients.
[210]12943_2020_1201_MOESM2_ESM.pdf^ (252.5KB, pdf)
Additional file 2. Supplementary materials and methods.
[211]12943_2020_1201_MOESM3_ESM.pdf^ (1.6MB, pdf)
Additional file 3 Figure S1. The expression and correlation analysis
between ZFAS1 and snoRNP complex based on the TCGA database. Figure S2.
The (m) RNA expression of ZFAS1, NOP58, SNORD12C, and SNORD78 in CRC
tissues and matched tumor-adjacent controls (n = 30). Figure S3. The
correlation analysis of ZFAS1, NOP58, SNORD12C/78 expression in 30
paired CRC and control tissues. Figure S4. The effect of ZFAS1 on cell
proliferation and apoptosis in CRC cells. Figure S5. The correlation of
ZFAS1 and NOP58 expression and prognosis evaluation in CRC patient
tissues. Figure S6. The 2′-O-Me activity levels after interfering ZFAS1
and/or NOP58 expression. Figure S7. The 2′-O-Me activities mediated by
SNORD12C and SNORD78 after silencing SNORD12C and SNORD78.
Acknowledgements