NADcapPro and circNC: methods for accurate profiling of NAD and non-canonical RNA caps in eukaryotes

Abstract

Accurate identification of NAD-capped RNAs is essential for delineating their generation and biological function. Previous transcriptome-wide methods used to classify NAD-capped RNAs in eukaryotes contain inherent limitations that have hindered the accurate identification of NAD caps from eukaryotic RNAs. In this study, we introduce two orthogonal methods to identify NAD-capped RNAs more precisely. The first, NADcapPro, uses copper-free click chemistry and the second is an intramolecular ligation-based RNA circularization, circNC. Together, these methods resolve the limitations of previous methods and allowed us to discover unforeseen features of NAD-capped RNAs in budding yeast. Contrary to previous reports, we find that 1) cellular NAD-RNAs can be full-length and polyadenylated transcripts, 2) transcription start sites for NAD-capped and canonical m 7 G-capped RNAs can be different , and 3) NAD caps can be added subsequent to transcription initiation. Moreover, we uncovered a dichotomy of NAD-RNAs in translation where they are detected with mito-chondrial ribosomes but minimally on cytoplasmic ribosomes indicating their propensity to be translated in mitochondria.

Full text

ARTICLE
NADcapPro and circNC: methods for accurate
profiling of NAD and non-canonical RNA caps in
eukaryotes
Sunny Sharma1✉, Jun Yang1, John Favate 2, Premal Shah 2& Megerditch Kiledjian 1✉
Accurate identification of NAD-capped RNAs is essential for delineating their generation and
biological function. Previous transcriptome-wide methods used to classify NAD-capped
RNAs in eukaryotes contain inherent limitations that have hindered the accurate identifica-
tion of NAD caps from eukaryotic RNAs. In this study, we introduce two orthogonal methods
to identify NAD-capped RNAs more precisely. The first, NADcapPro, uses copper-free click
chemistry and the second is an intramolecular ligation-based RNA circularization, circNC.
Together, these methods resolve the limitations of previous methods and allowed us to
discover unforeseen features of NAD-capped RNAs in budding yeast. Contrary to previous
reports, we find that 1) cellular NAD-RNAs can be full-length and polyadenylated transcripts,
2) transcription start sites for NAD-capped and canonical m7G-capped RNAs can be dif-
ferent, and 3) NAD caps can be added subsequent to transcription initiation. Moreover, we
uncovered a dichotomy of NAD-RNAs in translation where they are detected with mito-
chondrial ribosomes but minimally on cytoplasmic ribosomes indicating their propensity to
be translated in mitochondria.
https://doi.org/10.1038/s42003-023-04774-6 OPEN
1Department of Cell Biology and Neurosciences, Rutgers University, Piscataway, NJ, USA. 2Department of Genetics, Rutgers University, Piscataway, NJ, USA.
✉email: ssharma@dls.rutgers.edu;kiledjian@biology.rutgers.edu
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The presence of modified nucleosides is a characteristic
feature of the majority of cellular RNAs. Modified
nucleosides generally ascend from the chemical modifica-
tion of a genetically encoded nucleoside. One exception is the
addition of an N7-methylguanosine to the 5ʹend of select
eukaryotic RNAs transcribed by RNA polymerase II referred to as
“m7G cap”1. Historically only eukaryotic transcripts were believed
to contain 5ʹend modifications since the original analyses of
bacterial RNA composition did not detect modification on the 5′
end2,3. However, seminal work in the late 1970s4and early 20005
established that nucleotide metabolites, by virtue of their ade-
nosine nucleotide moieties (NAD and FAD), could be utilized by
E. coli and T7 RNA polymerases as initiating nucleotides for RNA
synthesis in vitro. The initial evidence that NAD caps are indeed
incorporated into RNA in cells was provided by mass spectro-
metry (LC-MS) in bacteria6. Nonetheless, due to the inability of
MS analysis to provide the sequence context, the physiological
role of these noncanonical caps remained elusive.
To assess the significance of the presence of the NAD cap in
cellular RNA, a chemo-enzymatic and Next Generation Sequen-
cing (NGS) based method, known as NAD captureSeq (NAD-
capSeq), was introduced in 20157. This method not only
corroborated the presence of NAD-capped RNAs (hereafter
referred to as NAD-RNA) in bacteria but also divulged the
identity of the transcripts. NADcapSeq transpired to be a decisive
advance in assessing the function of these caps in RNA meta-
bolism. Subsequent NADcapSeq analyses of yeast8, plant9,
human10, and archaeal RNA11,12 have led to the identification of
a significant number of transcripts with a NAD cap and have
established the presence of the NAD cap as a 5′non-canonical
cap in all three domains of life.
NADcapSeq leverages the unique feature of Adenosine
Diphosphate Ribosyl Cyclase (ADPRC) from Aplysia californica
to replace nicotinamide with alkynyl alcohol, in a transglycosy-
lation reaction13. The ‘clickable’transglycosylation product is
next biotinylated by a copper-catalyzed azide-alkyne cycloaddi-
tion (CuAAC)7. After enriching the biotin-linked RNAs using
streptavidin beads, the RNAs are subjected to NGS-based RNA
Seq analysis to identify the NAD-RNAs. Although NADcapSeq is
a robust method for cataloging NAD-RNAs, recent studies have
highlighted two of its key limitations; (a) the dependence on the
use of copper ions as a metal catalyst for the Huisgen cycloaddi-
tion reaction14, and (b) ADPRC promiscuity15,16. Metal ions,
including copper, can induce RNA fragmentation, which could
hinder the detection of low-abundant RNAs, and manifest a bias
toward shorter fragmented reads from the 5′termini of relatively
abundant RNA17. A further limitation of profiling NAD-capped
RNAs by NADcapSeq in eukaryotes involves the residual pro-
miscuity of the ADPRC enzyme on the m7G-capped RNAs16
(hereafter referred to as m7G-RNA). Depletion of m7G-RNA by
an anti-m7G cap antibody has been used as one strategy to
minimize the level of m7G-RNA in the NAD-RNA population16.
Although this is an improvement, additional approaches to
streamline NAD cap detection are indispensable for the rapid and
accurate profiling of the NAD-RNAs in eukaryotes. Furthermore,
the current lack of high-resolution mapping of the NAD position
in eukaryotes precludes mechanistic determination of NAD cap
addition.
Here we report two methods, NAD cap profiling (NADcapPro)
Seq, and intramolecular circularization of Noncanonical-Capped
(circNC) RNA. NADcapPro Seq builds on the recently reported
use of copper-free Strain-Promoted Azide-Alkyne Cycloaddition
(SPAAC)14 to identify NAD-capped transcripts in Arabidopsis16,
which depended on m7G cap antibody depletion to minimize
contaminating m7G-RNAs. NADcapPro Seq circumvents the
antibody depletion by coupling the robust m7G decapping
activity of Dcs118 prior to the SPAAC reaction. Dcs1 abolishes
the unfavorable residual enzymatic activity of ADPRC towards
canonical m7G cap. Moreover, direct visualization of the NAD-
capped RNAs was possible with streptavidin-conjugated IRDye.
A second method, circNC, provides an independent mechanism
to detect non-canonical capped RNAs at the nucleotide level of
precision. Analysis of NAD-capped RNAs initially detected by
NADcapPro subjected to circNC enabled us to extrapolate that
NAD caps are not only incorporated as noncanonical initiating
nucleotides in place of ATP but, can also be post-transcriptionally
added in eukaryotic cells. Furthermore, NAD caps were found to
be associated with mitochondrial ribosomes, raising the possibi-
lity that they can be translated in mitochondria.
Results
Development of NADcapPro and visualization of NAD-RNAs.
A key limitation of the NADcapSeq method used previously for
detecting NAD-RNAs is the use of copper ions as a metal catalyst
for the Huisgen cycloaddition reaction14. An alternative approach
based on copper-free chemistry, the strain-promoted azide-alkyne
cycloaddition (SPAAC) reaction19 was recently reported16.
Instead of using 4-pentyn-1-ol (NAD capSeq), 3-azido-1-
propanol is used for the ADPRC-catalyzed transglycosylation
reaction (Fig. 1a) where an azide moiety instead of an alkyne is
generated to replace nicotinamide, which would then become
amenable to the SPAAC reaction19. The azide residue introduced
upon transglycosylation can be subjected to the autocatalytic
alkyne moiety of biotin-PEG4-Dibenzylcyclooctyne (biotin-
PEG4-DBCO) to obviate the need of using copper ions (Fig. 1a
and Supplementary Fig. 1a).
We expanded on the use of the previously reported SPAAC
approach16 with two important modifications including direct
visualization of the NAD-RNAs and elimination of contaminat-
ing m7G-capped RNAs. In vitro transcribed NAD-RNAs and
triphosphate (pppA) RNAs were subjected to the SPAAC reaction
and the biotin-conjugated RNA (corresponding to NAD-RNA);
was visualized by near-infrared (IR) IRDye®800CW streptavidin-
based detection. As observed in Fig. 1b, IRDye®800CW
streptavidin enables direct visualization of biotinylated NAD-
RNAs and importantly validated the feasibility of the SPAAC
reaction for detecting NAD-RNAs. This was further verified by
using the same NAD-RNA substrate subjected to NAD decapping
(deNADding) by Rai1 treatment prior to the SPAAC reaction
which eliminated the signal (Supplementary Fig. 1b). Rai1 is a
well-characterized deNADding enzyme that removes the intact
NAD moiety without degrading the subsequent RNA10. More-
over, the sensitivity of IR dye for the detection of NAD-RNAs is
in the fmol range. SPAAC reactions with different concentrations
of in vitro transcribed NAD-RNAs demonstrated that biotin-
conjugated RNAs can be detected by the IR dye at levels as low as
15.6 fmol (Supplementary Fig. 1c). Importantly as previously
reported20, negligible levels of background autofluorescence and a
higher signal-to-noise ratio is apparent for the IR dye approach.
Next, to assess the advantage of SPAAC over CuAAC7in terms of
preserving the RNA integrity, total RNA isolated from yeast cells
was subjected independently to SPAAC and CuAAC reactions. As
expected, using previously published conditions for the CuAAC
reactions led to RNA degradation while the SPAAC reactions
minimally compromised RNA integrity (Fig. 1c). However,
whether further optimization of the CuAAC reaction may also
minimize RNA degradation is not clear.
Current approaches to scoring NAD-capped RNAs in eukaryotes
are limited by the detection of background residual m7G-capped
RNA10,16 which conflates the two populations of caps. To begin
optimizing eukaryotic NAD-RNA isolation, we delineated the
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nature of this background more systematically by assessing the
residual activity of ADPRC on canonical-capped RNAs, especially
m7GpppA and m7GpppG (account for the majority of transcrip-
tion start site (TSS) base21). Consistent with previous reports10,16,
detection by IR dye of biotinylated RNA analyzed by SPAAC
revealed residual activity on m7GpppA capped RNAs (Fig. 2a, and
Supplementary Fig. 7). Interestingly, this was not observed with
m7GpppG RNAs demonstrating a level of specificity previously
unknown for adenosine as the first transcribed nucleotide. With the
recent mass spectrometry-based estimates of relatively higher
m7GpppA-capped RNA to NAD-RNA9even minor levels of
detection could skew NAD cap detection.
To profile bona fide NAD-RNAs more accurately in eukar-
yotes, we developed a modified SPAAC approach to enzymati-
cally remove m7G-RNAs from the reaction. We leveraged the
selective activity of the yeast scavenger decapping enzyme Dcs118
and its robust pyrophosphatase activity on m7G-capped RNA
that hydrolyzes the phosphodiester bond between the gamma and
beta phosphates to release a 5′-diphosphate-RNA (ppRNA) and a
7-methyl-guanosine monophosphate18 prior to SPAAC modifi-
cation. We termed this approach NAD cap profiling
(NADcapPro).
To assess the feasibility of NADcapPro, a mixture of in vitro
transcribed 75 nts m7GpppA-capped RNA and 40 nts NAD-RNA
Fig. 1 SPAAC chemistry is superior to CuAAC for characterizing NAD-capped RNAs. a Schematic illustration of the SPAAC reaction to profile NAD-
capped RNAs. bTen picomoles of 40 nts in vitro transcribed RNA with a 5′triphosphate (pppA) or 5′NAD cap were subjected to the SPAAC reaction. The
RNAs were resolved on an agarose gel at the indicated concentration either without or with the addition of 20 µg of background total yeast RNA and
transferred onto a Nitrocellulose membrane. The biotin-conjugated RNAs (NAD) were visualized by near-infrared (IR) IRDye®800CW streptavidin-based
detection using Odyssey Fc (Li-Cor Biosciences). The NAD cap or uncapped RNA is represented by the NAD of pppA followed by a line that denotes the
RNA. cTwenty micrograms of total yeast RNA were subjected to SPAAC and CuAAC reactions and the RNAs were detected using IRDye®800CW. The
CuAAC reactions led to RNA degradation while the SPAAC reactions minimally affect the integrity of the RNA.
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at molar ratios of ~50:1 was used in the absence or presence of
Dcs1 treatment. Visualization of the RNA following the SPAAC
reaction was carried out with DBCO-Cy3 to visualize the clicked
RNAs, avoiding the prerequisite to blot the RNA onto a
membrane. As shown in Fig. 2b, ADPRC indeed displayed
a noticeable activity towards m7GpppA-capped RNA in the
absence of Dcs1 treatment with robust detection of the m7GpppA-
capped RNA. In contrast, treating the RNA mixture with
Dsc1 specifically abolished the reactivity of the ADPRC towards
the m7GpppA capped RNA without affecting the detection of
NAD-RNAs, demonstrating the feasibility of NADcapPro for
identifying NAD-RNAs in eukaryotes. This was next validated by
using endogenous polyA+RNAs from WT cells treated with and
without Dcs1 prior to the SPAAC reaction (Fig. 2c). Visualization
of the RNAs revealed a substantial reduction in IR dye signal in
the NADcapPro lanes compared to the SPAAC lanes, consistent
with the selective removal of m7G RNA detection (Fig. 2d). In
addition, the RNA subjected to NADcapPro can be retained and
eluted from streptavidin beads (Fig. 2e) to enable subsequent
analysis.
Transcriptome-wide mapping of NAD-RNAs in yeast.To
demonstrate the feasibility of NADcapPro at the genomic scale,
we generated standard RNA-seq, SPAAC-seq and NADcapPro-
seq datasets derived from a WT strain and a strain disrupted for
deNADding activity of a prominent deNADding protein Xrn1
Fig. 2 ADPRC cross-reactivity with m7GpppA capped RNAs and NADcapPro as a superior method to resolve the inherent limitations of previous
methods. a Ten picomoles of 40nts in vitro transcribed NAD-, m7GpppA- and m7GpppG-capped RNA were subjected to the SPAAC reaction and
visualized using IRDye®800CW. Detection of biotinylated RNA analyzed by SPAAC revealed residual activity only on m7GpppA-capped RNA. bIn vitro
transcribed m7GpppA-RNA (75nts) with and without Dcs1 and 40nts of NAD-capped RNA were mixed at molar ratios of (50:1) in a SPAAC reaction. The
RNAs were visualized directly in the gel. ADPRC exhibited a noticeable activity towards m7GpppA capped RNA in the absence of Dcs1 treatment.
Treatment of the RNA mixture with Dcs1 explicitly abolished the reactivity of the APRC towards the m7GpppA-capped RNA without compromising the
detection of NAD-RNA. cSchematic illustration of NADcapPro protocol used in the present study. dComparison of the SPAAC and NADcapPro reactions
using endogenous polyA+RNA isolated from yeast were resolved by agarose gel electrophoresis and visualized using IRDye®800CW. A substantial
reduction in NAD-RNA signal is apparent in the NADcapPro lanes compared to the SPAAC lanes, consistent with the selective removal of m7G-capped
RNA detection. ePolyA+RNA (15 µg) was subjected to SPAAC and NADcapPro. The affinity-purified biotinylated RNA corresponding to NAD-capped
RNA was eluted from streptavidin beads and visualized using IRDye®800CW as in (d).
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(xrn1-H41A)22. Differential expression analyses using DESeq223
identified transcripts enriched in SPAAC or NADcapPro relative
to standard RNA-seq. We classified a gene as potentially capped if
it was enriched by at least twofold over the standard RNA-seq at a
q-value ≤0.01 (Fig. 3a). Based on these two metrics, SPAAC-
NAD-seq identified 268 transcripts, whereas NADcapPro seq
identified 769 NAD-capped transcripts in WT cells and 830
NAD-capped transcripts in the xrn1-H41A cells. The xrn1-H41A
protein is deNADding deficient yet retains the 5′-3′exoribonu-
clease activity22 and is expected to have a higher number of
NAD-RNAs. However, the differential is modest since yeast
harbor additional deNADding enzymes10,17 and Xrn1 appears to
Fig. 3 NADcapPro Seq. a The relationship between RNA abundance and fold-change in the experimental condition is indicated at the top of each panel.
The x-axis indicates RNA abundance from RNAseq derived from WT (blues) or xrn1 mutant (oranges) cells. The y-axis indicates the DESeq2 fold change.
Colored points indicate a fold-change log
2
≥1 and q-value ≤0.01. The number in the upper right indicates the number of genes meeting these criteria.
bNADcapPro can capture low abundance transcripts. The distribution of average TPMs for the colored genes in (a) is shown. ****p< 0.0001 for a one-
sided t-test testing if the NADcapPro distributions are less than the SPAAC distributions. Venn diagram overlap of the indicated gene sets in WT,
NADcapPro and Xrn1-targeted NAD-RNAs are represented in (c) and (d) respectively. Gene ontology categories for the indicated gene sets in WT,
NADcapPro (e) and Xrn1-targeted NAD-RNAs (f). All source data are contained in the GSE217259 repository.
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have a restricted NAD-RNA target population22. In addition,
although a higher number of NAD-RNAs evident with NAD-
capPro relative to SPAAC may appear counterintuitive, it is the
removal of the abundant contaminant m7G-capped transcripts
that enables the detection of these low-abundance NAD-RNAs.
For example, four transcripts that were scored below the sig-
nificance threshold in SPAAC (RPL21B, ENB1, IMD4, and
SGF11) were identified as NAD-RNAs by NADcapPro (Fig. 3a)
and further validated to be NAD-capped by an independent
approach (see below).
Additional support for the specificity of NADcapPro was
evident upon analysis of the identified transcript’s abundance.
Whereas SPAAC is only able to detect highly expressed NAD-
RNAs, NADcapPro can detect NAD-RNAs with lower expres-
sion. This is illustrated in Fig. 3b, where the Transcripts Per
Kilobase Million (TPM) distribution of potentially capped genes
from NADcapPro are lower than the SPAAC distribution.
Interestingly as shown in Fig. 3c, only 63 transcripts (~8% of
the total NAD-capped transcripts) overlapped between SPAAC
and NADcapPro seq, a strong indication that most transcripts
identified by SPAAC seq corresponded to the relatively higher
abundance of m7GpppA-capped transcripts which in turn likely
impeded the capture of the moderately abundant NAD
transcripts. A comparison of the number of NAD transcripts
responsive to the robust Xrn1 deNADding activity (xrn1-H41A)
to that of WT cells revealed 346 transcripts that were directly
responsive to Xrn1 deNADding (Fig. 3d) while the remaining
transcripts appear to be regulated by deNADding enzymes other
than Xrn110,17. This is in contrast to our initial use of the CuAAC
approach to identify NAD transcripts responsive to Xrn1 that
only identified the abundant mitochondrial NAD transcripts with
confidence22. Collectively, these results demonstrate that NAD-
capPro seq is more robust and adept at capturing high and low-
abundance NAD transcripts that are otherwise not detected with
the standard NAD-RNA isolation approaches.
Gene Ontology (GO) term analysis was carried out to ascertain
the potential biological processes the refined list of NAD-RNAs
may be involved in (Figs. 3e and 3f). Investigation of the 706
transcripts and 346 transcripts from the WT and xrn1-H41A
mutant along with the transcripts in common to both
(Supplementary Fig. 2) respectively yielded a diverse array of
pathways. In addition to the chemical and transcriptional
machinery of RNA pol II, a high cluster frequency was observed
for nuclear-encoded mitochondrial genes involved in both
mitochondrial organization and mitochondrial translation in
the WT pool. This is consistent with NAD being an important
cofactor for RNA transcription and mitochondrial function. The
Xrn1-responsive NAD-RNAs also were similarly involved in
chemical and transcriptional responses, in addition to transmem-
brane transport and the meiotic cell cycle. The most striking GO
term among the top pathways that exhibited a relatively greater
cluster frequency was “Transposition”encompassing the process
involved in mediating the movement of discrete segments of
DNA between nonhomologous sites. Intriguingly, these tran-
scripts were undetected with standard RNA sequencing but are
enriched by more than log
2
> 10 by NADcapPro (highlighted in
red in Fig. 3a) suggesting that transcripts belonging to this GO
term are predominantly NAD-capped. The importance for NAD
capping of these transcripts (Supplementary Table 1) remains to
be determined; however, it is tempting to speculate that NAD
capping of these RNA may be a major contributor to their low
abundance consistent with NAD caps promoting RNA decay10.
Non-canonical caps can be added following transcription
initiation. The current dogma postulates that NAD caps are
incorporated during transcription initiation as noncanonical
nucleotides in place of ATP as the first transcribed nucleotide24,25.
Although there is in vitro evidence in support of this hypothesis,
cell-based evidence is lacking in eukaryotes. To directly address
whether NAD caps can be added by a transcription initiation
independent mechanism we developed an approach to map the
precise NAD metabolite nucleotide addition site within transcripts
validated by NADcapPro to possess NAD caps. We built on two
previously reported methodologies, CapZymeSeq26 and RNA end
circularization27 to develop an orthogonal approach which we
termed intramolecular circularization of Noncanonical-Capped
RNA, circNC (Fig. 4a). Here we leveraged the robust in vitro
deNADding activity of Rai1, which removes the intact NAD
moiety from the 5′end of an RNA to generate a 5′monopho-
sphate RNA10. Importantly, Rai1 does not hydrolyze m7G-capped
transcripts10 (Supplementary Fig. 3a). The resulting monopho-
sphate RNA is intramolecularly circularized using T4 RNA ligase
to produce a circular RNA with the 5′untranslated regions (UTR)
fused to the 3′end polyA tail followed by the 3′UTR region. The
use of transcript-specific primers enables cDNA synthesis followed
by specific PCR amplification across the junction.
CircNC was used to validate four of the nuclear-encoded NAD
transcripts identified by NADcapPro from Fig. 3a: RPL21B,
ENB1, IMD4, and SGF11 (Supplementary Fig. 4a). The use of
circNC on these yeast NAD-capped mRNAs yielded the expected
PCR product following Rai1 deNADding (Fig. 4b). The size of the
band was comparable to that generated when the RNA was
decapped with the m7G-capped mRNA decapping enzyme
(MDE) (Fig. 4b) which is unable to hydrolyze NAD-capped
RNA (Supplementary Fig. 3a) or triphosphate RNA (Supplemen-
tary Fig. 3b). To verify the specific amplicons and assess the
sequence context of the respective NAD caps, the PCR products
were subjected to next-generation sequencing (Supplementary
Fig. 4b). The sequence analyses of the Rai1 and MDE-treated
samples are shown in Fig. 4c and Supplementary Data 1. Since
transcription initiation sites can be variable, we focused on the
most prominent transcripts for each cap and each mRNA. Two
important differences are apparent when comparing the canoni-
cal m7G and NAD transcripts. First, the predominant 5′ends of
the m7G-capped and NAD-capped transcripts are distinct and
they do not share a common transcription start site. Second, two
of the four NAD transcripts analyzed do not start with an
adenosine moiety in the coding strand. The importance of this
latter observation is that non-canonical caps are believed to be
incorporated onto the 5′ends of prokaryotic mRNA by the
incorporation of NAD rather than ATP as the first transcribed
nucleotide during transcription initiation24. By extension, this has
also been presumed to be true in eukaryotic cells. Such a
mechanism necessitates the presence of an adenosine nucleotide
in the coding strand of the DNA. However, as apparent with the
NAD cap positions in ENB1 and SGF11, corresponding
adenosine is not evident at the NAD cap addition site in the
reference coding sequence. In the case of RPL21B and IMD4, the
position of the NAD does contain an A residue consistent with a
co-transcriptional NAD addition. We conclude NAD cap
addition can be carried out by a post-transcriptional initiation
process of NAD capping. Furthermore, the analyzed transcripts
were all polyadenylated, indicating that NAD transcripts are
intact full-length mRNAs, contrary to previous reports17. The 5′
end skewed representation of NAD transcripts in previous reports
was likely a byproduct of CuAAC-mediated RNA fragmentation
during the NAD capture.
Nuclear-encoded NAD-RNAs are likely not translated. Since
nuclear encoded NAD-RNAs have intact 5′and 3′UTR and are
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polyadenylated akin to canonical capped transcripts, we next
tested if the NAD transcripts are translated. Initial reports using a
transfected NAD-capped RNA in mammalian cells did not detect
any appreciable level of translation10. However, definitive inter-
pretations of these data were complicated by the fact that an
exogenous transcript was used rather than an endogenous NAD-
RNA. Support for the translation of NAD-RNAs was subse-
quently reported in Arabidopsis thaliana where NAD-capped
transcripts were detected within cytoplasmic polysomes9. How-
ever, as these studies relied on the NADcapSeq approach, it was
not clear whether the detected RNAs were indeed NAD-capped
as previously reported16.
To assess whether NAD-RNA can associate with actively
translating pools of RNA, we isolated RNAs from polysome
gradients and subjected them to NADcapPro (Fig. 5a). PolyA+
RNA was isolated from the free pool, monosome, and merged
polysome fractions and analyzed by NADcapPro. NAD-RNAs
were visualized using IR dye. Intriguingly, NAD-RNAs were
predominantly detected in the free pool representing the fraction
of cellular polyA+RNAs that are not engaged in translation and
minimally in the translationally active polysome fractions (Fig. 5b
and Supplementary Fig. 8). To further validate this observation,
polyA+RNAs isolated from the same fractions were subjected to
circNC analysis. As shown in Fig. 5c, circNC of two NAD
transcripts, RPL21B and SGF11, further corroborated the
NADcapPro results and revealed that most NAD transcripts are
excluded from translationally active ribosomes and are likely not
engaged in translation.
Fig. 4 circNC as an orthogonal method for non-canonical-capped RNA detection. a Schematic illustration of the circNC method. bTranscript-specific PCR
of validated NAD-capped RNAs subjected to circularization following Rai1 treatment (NAD-RNA) or MDE treatment (m7G-RNA) as illustrated in (a) and
resolved on 2% TAE-agarose gel. cSequence alignment of the Rai1 and MDE-treated samples derived from next-generation sequencing of the PCR
products generated from the nuclear-encoded transcripts are shown. The alignment represents the most prevalent annotated reads spanning the 5′-3′
junction. The percentage of reads corresponding to the depicted sequences are shown on the right of each sequence. The green color of the reference
sequence represents the 5′UTR while the red region denotes the open reading frame starting with the ATG initiation methionine sequence.
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Mitochondrial NAD-RNAs are associated with translating
ribosomes. The initial report identifying NAD-capped transcripts
in S.cerevisiae indicated a predominance of NAD caps on mito-
chondrial encoded transcripts8, and two, Cox2 and 21 S rRNA,
were subsequently validated by an independent method22,24.
Yeast mitochondria encode 8 major protein-coding genes –Cox1,
Cox2, Cox3, COB, SCE1, ATP6, ATP9, and VAR1 along with two
mitochondrial ribosomal RNAs -15 S and 21 S rRNA, and 24
tRNAs. Since mitochondrial transcripts lack a polyA tail, they
were not included in our above NADcapPro analysis. We next set
out to establish a comprehensive NAD-RNA profile of protein-
coding and rRNA yeast mitochondrial transcripts by SPAAC
analysis of total RNA isolated from purified mitochondria and
assessed NAD transcripts by Northern blotting using the RNA
eluates from the affinity purified NAD-RNAs. The lack of m7G
caps on mitochondrial transcripts precluded the need to employ
Fig. 5 Nuclear-encoded NAD-capped RNAs are not translated. Polysome gradient profile of WT cells. Sucrose-density gradient ultracentrifugation was
used to fractionate the free pool, ribosomal subunits, monosome, and polysomes. aOptical density profiles (Abs
260nm
) of polysome gradient are shown.
PolyA+RNA was isolated from the free pool, monosome, and merged polysome fractions and analyzed by NADcapPro. bNAD-RNAs were visualized
using IR dye. Noticeably, NAD-capped RNAs were exclusively detected in the free pool representative of the fraction of polyA+RNAs not engaged in
translation. cPolyA+RNA from the polysome gradients was subjected to circNC analysis and transcript-specific PCR of the RPL21B and SGF11 circularised
RNAs are shown. The Rai1 and MDE treated lanes represent NAD- and m7G-capped RNAs respectively. The Ctrl is RNA that was not treated with either
Rai1 or MDE enzymes. PCR products were resolved on a 1.5% TAE-agarose gel.
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NADcapPro. Only 5 NAD mitochondrially encoded transcripts
were detected to be NAD-capped (Fig. 6a, and Supplementary
Fig. 9). Three corresponded to protein-coding transcripts –Cox1,
Cox2, and ATP9, and two were the non-coding mitochondrial
ribosomal RNAs –21 S rRNA and 15 S rRNA (Fig. 6a, and
Supplementary Fig. 9). We further validated NAD capping of
ATP9, Cox1, and 15 S rRNA using DNAzymes (Supplementary
Fig. 5, and Supplementary Fig. 8).
The presence of protein-coding mitochondrial mRNAs posses-
sing a NAD cap prompted us to test whether these transcripts are
found in actively translating polysomes. To address this question,
mitochondrial ribosomes were affinity purified using a recently
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established protocol28. A FLAG-tagged nuclear-encoded mito-
chondrial small subunit ribosomal protein MRPS17 was used as
bait to affinity purify mitochondrial ribosomes (Fig. 6c, d, and
Supplementary Fig. 10) under conditions that optimize immu-
noprecipitation of membrane-associated mitoribosomes whilst
retaining subunit association28 (Fig. 6b). Northern Blot analysis
of the immunopurified products demonstrated that intact
mitoribosomes consisting of both 15 S and 21 S rRNA subunits
as well as an mRNA (Cox2) are affinity-purified (Fig. 6e),
consistent with actively translating ribosomes as previously
reported28. To assess the presence of NAD transcripts in this
translationally active ribosome pool, boronate affinity
electrophoresis29,30 was used to analyze two of the protein-
coding transcripts, Cox2 and ATP9, and two non-coding rRNA
(15 S and 21 S) in more detail. This method renders direct
visualization of both NAD-capped and uncapped RNA popula-
tions by specifically impeding the mobility of NAD-RNAs in the
gel due to the transient formation of diesters between
immobilized boronic acid and the ribose moiety29,30.
DNAzyme-mediated RNA cleavage was used to generate 5′-end
containing sub-fragments of defined length22, and a Northern
blot analysis with the 32P labeled transcript-specific probes was
used to corroborate the presence of NAD-capped transcripts.
Remarkably, unlike nuclear-encoded transcripts, both ATP9 and
Cox2 NAD mRNAs are associated with the translationally active
mitochondrial ribosomes (Fig. 6f, and supplementary fig. 11).
Interestingly, the Cox2 mRNA also possesses NADH-capped
RNA as previously reported24, while this was not detected on any
of the other mitochondrial transcripts. The rationale of this
distinction is not clear. An additional intriguing observation was
that only one of the NAD-capped derivatives of the ribosomal
RNAs (15 S rRNA) is detected in the mature ribosome, but not
the 21 S rRNA (Fig. 6g, and supplementary fig. 11) even though
both can be NAD-capped (Fig. 6a). Collectively, our data
demonstrate that, unlike cytoplasmic NAD-RNAs, NAD-capped
mitochondrial mRNAs are associated with translating ribosomes
and may be translated.
Discussion
The presence of NAD as a 5′noncanonical cap in cellular RNAs
has been established in various organisms7,8,10,16. To understand
the biological function of 5′NAD caps in cellular physiology, the
identification of cellular NAD-RNAs is imperative. The original
method introduced to catalog transcriptome-wide NAD-RNAs in
different organisms-NADcapSeq was instrumental in under-
standing the nature of NAD caps in prokaryotic RNA despite the
limitation of using copper-based click reaction (CuAAC) that
favored 5′RNA fragments. However, more pertinently, its
extrapolation into analyzing eukaryotic mRNAs with m7G caps
was problematic due to the promiscuity of the ADPRC reaction.
To address this issue, Hu et al16 reported the use of copper-free
SPAAC-mediated NAD-RNA isolation and proposed the use of
m7G-capped RNA depletion to enrich for NAD-RNA. Herein we
expanded on the use of copper-free SPAAC-mediated NAD-RNA
isolation and coupled the SPAAC reaction with a more robust
enzymatic approach to eliminate m7G-capped RNAs to accu-
rately profile NAD-RNAs in eukaryotes and further leveraged the
highly sensitive IRDye®800CW streptavidin-based detection to
visualize NAD-RNAs at concentrations as low as 15 fmol. Of
note, a recent report also used an enzymatic approach to mini-
mize m7G capped RNA within the NAD population by eluting
biotin-conjugated NAD-RNAs with a NAD decapping enzyme31.
NADcapPro incorporates the capacity of ADPRC to use an
azide for the transglycosylation reaction through a copper-free
click-chemistry-based SPAAC reaction that maintains RNA
integrity. It also addresses the cross-reactivity of ADPRC with the
canonical m7GpppA-caps in eukaryotes by exploiting the robust
m7G decapping activity of yeast Dcs118. Dcs1 treatment of RNA
prior to the ADPRC reaction eliminates ADPRC reactivity
towards the m7GpppA-capped RNAs (Fig. 2b) and enables affi-
nity purification of true NAD-RNAs along with the identification
of a relatively higher number of NAD-RNAs that previously
remained obscure. Importantly, we also provide an orthogonal
circNC method to validate and expand on the NADcapPro
approach. CircNC is a sensitive approach to identifying RNAs
harboring a non-canonical cap and enables precision mapping of
the cap addition site to the nucleotide level as well as the precise
3′termini of the RNA. While NAD caps are the primary focus of
our study, it is worth noting that Rai1 can also decap FAD32 and
dephospho CoA caps32. Therefore, although the RNAs used in
the circNC analysis consisted of RNAs validated by NADcapPro
to possess an NAD cap, the formal possibility remains that a
mixture of non-canonical caps may be detected. Future optimi-
zation with Rai1 mutants that can distinguish between distinct
non-canonical caps will further enhance the methodology.
Our approaches to NAD-RNA isolation and visualization
enabled a refined analysis of the NAD nucleotide metabolite-
capped class of RNAs. First, the length and state of NAD-RNAs
have been confounded by different results of whether NAD-
RNAs can be full-length or not10,17. circNC by virtue of detecting
the polyA tail at the junction of the circularized RNA enables an
assessment that NAD-RNAs can be full-length (Fig. 4a, c).
Second, the 5′ends of m7G-capped RNAs and their corre-
sponding NAD-capped transcripts were different and did not
map to the same start sites. Notably, although they did contain
relatively similar polyA tail lengths, two of the four NAD-capped
transcripts analyzed, SGF11 and IMD4, contained relatively
longer 3ʹUTRs than their m7G-capped counterpart transcripts
(Supplementary Data 1). Third, a comparison of the NAD cap
addition site to the genomic sequence revealed NAD caps that did
not align with corresponding adenosine in the genomic sequence
indicating that the NAD moieties were not incorporated in place
of adenosine as a noncanonical nucleotide during transcription
initiation and strongly implicate the existence of a post-
transcription initiation capping mechanism.
The best-characterized mechanism of NAD cap addition cur-
rently is the ab initio addition of non-canonical caps including
NAD caps. This is primarily based on in vitro studies demon-
strating that the RNA polymerases from both prokaryotes and
eukaryotes can add a NAD cap in place of ATP at the 5′-end
Fig. 6 Mitochondrial NAD-RNAs are associated with a translationally active pool of mitoribosomes. Comprehensive NAD-capped RNA profile of yeast
mitochondrial transcripts by SPAAC analysis. aNorthern blot analysis of the RNA eluates from the affinity-purified NAD-capped RNA by SPAAC.
bSchematic illustration of the strategy used to isolate the translationally active pool of ribosomes as explained recently by28.cEluates from MRPS17-
FLAG- IP were loaded onto a 4–12% Bis-Tris gel and stained with SYPRO Ruby. The no FLAG affinity purification was used as a control. dWestern blot
analysis with an anti-FLAG-tag antibody for the validation of the presence of MRPS17-FLAG-tagged protein. eNorthern Blot analysis of RNA isolated from
the MRPS17-FLAG- IP eluates. Northern Blot analysis of DNAzyme-generated 5′-end-containing subfragments of the RNA isolated from the MRPS17-
FLAG- IP eluates resolved on 0.3% 3-acrylamidophenylboronic acid 10% PAGE gel and detected with the 32P labeled transcript-specific probes of the
indicated mRNAs in (f) and ribosomal RNAs in (g).
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during transcription initiation25. An alternative mode of post-
transcriptional addition of the NAD caps has also been proposed
based on the presence of NAD caps on small nucleolar RNAs
(snoRNAs) whose 5′ends are generated by exoribonucleases and
cannot undergo ab initio capping10. In the present study, we
expand on these results and demonstrate in addition to snoRNAs,
cytoplasmic mRNA can also be NAD-capped post-tran-
scriptionally suggesting the presence of dedicated capping
machinery at least in S.cerevisiae. The demonstration of both
human and plant snoRNAs with NAD caps10,33 also suggests that
post-transcriptional NAD capping is likely a general mechanism
in eukaryotes. Our data is consistent with two modes of NAD cap
addition, one being ab initio addition and the second through a
yet uncharacterized NAD capping mechanism. Collectively, our
findings reveal the complex nature of cellular NAD capping and
the limitation of our current understanding of this intriguing
modification.
The ability to more accurately identify NAD-RNAs enabled us
to assess their presence on actively translating ribosomes and
address whether NAD-RNAs can be translated. Consistent with
our previous assessment that in vitro generated exogenous NAD-
RNAs are not translated10 we demonstrated that nuclear-encoded
NAD-RNAs minimally interact with a translationally competitive
pool of cytoplasmic ribosomes and do not appear to be engaged
in translation in yeast cells under normal growth conditions.
Remarkably, a more dramatic outcome was detected in mito-
chondria. Mitochondrial protein-coding NAD-RNAs were
detected to be engaged within the translationally competent pool
of mitochondrial ribosomes, consistent with undergoing active
translation. One possibility for the differing states of translat-
ability between the cytoplasmic and mitochondrial transcripts
could be the requirement of the m7G cap for the cytoplasmic
translation1but not mitochondrial translation34. However, the
formal possibility that these NAD transcripts are not engaged in
translation but function as regulatory RNAs cannot be ruled out.
Future studies will address any potential role of NAD cap in
mitochondrial translation.
Methods
Yeast growth and media. Yeast cells (BY4741 strain background) WT and xrn1-
H41A22 were grown at 30 °C in (1% w/v yeast extract, 2% w/v peptone, 2% w/v
glucose/glycerol). All yeast strains used in the present study are listed in Supple-
mentary Table 2.
In vitro transcription of NAD and m7G-capped RNAs. RNAs containing NAD
and m7G cap structures were synthesized by in vitro transcription from synthetic
double-stranded DNA template ɸ2.5-NAD-40, ɸ2.5-NAD-75 containing the T7
ɸ2.5 promoter, and a single adenosine within the transcript positioned at the
transcription start site (Supplementary Table 3). For m7G-capped RNA, m7G-(5′)
PPP (5′)-A RNA Cap Structure Analog (New England Biolabs) was included in the
transcription reaction, whereas for NAD-RNA, NAD was used instead of ATP. In
vitro transcription was carried out at 37 °C overni ght, using HiScribeTM T7 High
yield RNA Synthesis kit (New England Biolabs (NEB)). Following in vitro tran-
scription, RNA was purified using Monarch®RNA Cleanup Columns (NEB) as per
the manufacturer’s instructions.
RNA in vitro deNADding and decapping assays. For deNADding, the NAD-
RNAs were incubated with 100 nM SpRai1 recombinant protein in NEB buffer
(100 mM NaCl, 50 mM Tris-HCl, 10 mM MgCl
2
, pH 7.9). Reactions were incu-
bated at 37 °C for 1 hour, and the deNADed RNA was purified using
Monarch®RNA Cleanup Columns (NEB) as per the manufacturer’s instructions.
For decapping of both in vitro transcribed m7G capped RNA and polyA
enriched RNA from yeast cells, yeast DcpS enzyme (yDcpS; NEB) which we
referred to by the SGD nomenclature, Dcs1, was used. Typically, the reaction was
performed in 50 µL volume using 4 units per µg of capped RNA following the
manufacturer’s protocol. The decapped RNA was purified using Monarch®RNA
Cleanup Columns (NEB) as per the manufacturer’s instructions.
Total RNA extraction and polyA+RNA enrichment. Total RNA from yeast
strains was isolated with the acidic hot phenol method35 and treated with DNase
(Promega) according to the manufacturer’s protocol. Total RNA from human cells
was isolated using TRIzol reagent (Thermo Fisher Scientific) according to the
manufacturer’s protocol and treated with DNase as explained above to eliminate
DNA. To enrich for poly-adenylated RNA, total RNA purified as above was used as
the input for the Poly(A) Purist MAG Kit (Thermo Fisher Scientific) following the
manufacturer’s protocol. We retrieved ~1.5 µg of polyA RNA from 100 µg of total
yeast RNA.
Cu(I)-catalyzed Azide–Alkyne Cycloaddition (CuAAC) and Strain-promoted
Alkyne-Azide Cycloaddition (SPAAC) reactions. To biotinylate NAD-capped
RNAs using CuAAC, 50 µg of total RNA was first incubated with 10 µL of 4-
pentyn-1-ol (Sigma-Aldrich), 10 µL of (125 µg/mL) of Adenosine diphosphate-
ribosylcyclase (ADPRC) in a 100 µL reaction containing 50 mM HEPES, 5 mM
MgCl
2
(pH 7.0), and 40 U of RNasin®Ribonuclease Inhibitor (Promega) at 37 °C
for 1 h. After the ADPRC treatment, RNAs were incubated with 250 µM biotin-
PEG3-azide, freshly mixed 1 mM CuSO
4
, 0.5 mM THPTA, 2 mM sodium ascorbate
in a 100 µL reaction with 50 mM HEPES, 5 mM MgCl
2
(pH 7.0), and 40 U of
RNasin®Ribonuclease Inhibitor (Promega) at 30 °C for 30 min. The RNA was then
precipitated with ethanol in the presence of 2 mM EDTA and 2 M ammonium
acetate10,36. For reactions using the SPAAC approach, ADPRC was first used to
replace the nicotinamide and add an azide residue onto the NAD-RNA. Typically,
these reactions were performed in a 100 µL volume with either total RNA or polyA-
purified RNA dissolved in DEPC water, 10% (v/v) of 3-azido-1-propanol (Sigma)
together with 10 µL of (125µg/mL) of ADPRC (Sigma) in ADPRC reaction buffer
(50 mM HEPES (pH 7.0) and 5 mM MgCl
2
). The reaction was carried out at 37 °C
for 1 hour. The transglycosylated RNA with the azide residues was next purified
using Monarch®RNA Cleanup Columns (NEB) and eluted in 20 µL of DEPC water.
For the SPAAC reaction, dibenzocyclooctyne-PEG4-biotin (DBCO-biotin, Sigma)
was used as the alkyne moiety for the cycloaddition. Following the ADPRC reac-
tion, both total RNA and polyA enriched RNA in pure water were mixed with 1x
volumes of MK-Gel Loading Buffer (90% Formamide, 15 mM EDTA, and 0.025%
SDS) and 500 μM DBCO-biotin (Sigma). Typically, these reactions were 20 μL MK-
Gel Loading Buffer, 18 μL RNA (from ADPRC reaction), and 2 μL 10 mM stock of
the DBCO reagent. The reactions were performed at 55 °C for 15 minutes to
denature the RNA and stoppe d by adding 310 μL water followed by purification of
the conjugated RNA using Monarch®RNA Cleanup Columns (NEB). The products
were analyzed by gel electrophoresis or subjected to affinity purification as
described below.
RNA gel electrophoresis, capillary blotting, and near-InfraRed fluorescent
imaging. For blotting analysis of either enriched RNA following biotin-mediated
affinity purification or directly after the SPAAC reaction, RNA was incubated at
90 °C for 2 min in formaldehyde loading buffer (FLB) (50% Formamide, 6% for-
maldehyde, 50 mM HEPES (pH 7.8), 0.5 µg/mL ethidium bromide and 10% gly-
cerol). Samples were then loaded onto a 1% agarose gel and resolved by
electrophoreses at 120 V for 1 h. Before tran sferring the gel onto a 0.45 μm
nitrocellulose membrane (Cytiva), total RNA was first visualized using a UV gel
transilluminator. RNA was transferred overnight by passive, upward transfer
facilitated by the capillary flow of the 10X SSC buffer. After transfer, RNA was
cross-linked to the NC using UV-C light (0.2 J/cm2). The membrane was next
blocked with Odyssey Blocking Buffer, PBS (Li-Cor Biosciences) for 30 minutes at
room temperature (RT). After blocking, IRDye®800CW streptavidin (Li-Cor
Biosciences) was diluted to 1:7,000 in Odyssey Blocking Buffer and stained the NC
membrane for 30 minutes at RT. The membrane was next washed three times with
PBST (ResearchProductInternational). Before scanning, the membranes were
briefly rinsed in 1x PBS and scanned by an Odyssey Fc (Li-Cor Biosciences) with
the software prearranged to auto-detect the signal intensity for both the 600 and
800 channels. The 600 channel was used for ethidium bromide and the 800 channel
was used for the detection of IRDye®800CW streptavidin.
To assess the residual activity of ADPRC towards the m7GpppA capped RNA,
we mimicked the in vivo differential by mixing in vitro transcribed 40 nts NAD-
RNA and 75 nts m7GpppA capped RNA at molar ratios of ~50:1. The mixture was
subjected to the SPAAC- reaction as discussed above with the only difference that
instead of DBCO-biotin, DBCO-Cy3 was used for cycloaddition. The RNA was
purified over the Monarch®RNA Cleanup Columns (NEB). The eluted RNA was
next mixed with an equal amount of 2X MK- Gel Loading Buffer and denatured at
65 °C before resolving on a 7 M 15% PAGE gel. The ethidium bromide-stained
RNA was first visualized in the gel using a UV gel transilluminator and the Cy3
conjugated RNA was detected using the Cy3 channel on a Typhoon scanner (GE)
NADcapPro seq. Fifteen micrograms of polyA RNA were isolated from three
independent biological replicates of both yeast WT and xrn1-H41 mutant and were
treated with Dcs1 as described above to remove m7G-caps. Samples without Dcs1
treatment (SPAAC) and ADPRC (minus ADPRC) were also processed similarly to
assess the contribution from m7G caps and any background noise due to highly
abundant RNAs. Following the SPAAC reactions, enrichment was achieved by
selective affinity to streptavidin beads10. Briefly, RNAs were incubated at 25 °C for
30 min with 15 μL of magnetic Dynabeads™MyOne™Streptavidin T1, which were
pre-blocked with 100 ng/µL of bacterial small RNAs in 100 μL of immobilization
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buffer containing 10 mM Tris-HCl (pH 7.5) 1 mM EDTA and 2 M NaCl. After
rigorous washing with wash buffer (10 mM Urea, 5 mM Tris-HCl [pH 7.5], 0.5 mM
EDTA, and 1 M NaCl) five times at 25 °C for 5 min each, biotinylated RNAs were
eluted by incubating the beads with 20 μL of MK-Gel Loading Buffer at 90 °C for
2 min. The biotinylated RNAs were next purified using Monarch®RNA Cleanup
Columns (NEB) as described above and eluted in 10 µL of nuclease-free water. The
samples were next flash-frozen in liquid nitrogen and were sequenced at GENE-
WIZ (Azenta Life Sciences).
NADcapPro sequencing analysis. Raw reads were processed for quality control
and adapter removal by fastp37. Following this, in silico rRNA depletion was
accomplished by using hisat238 to align the reads to rRNA sequences and save the
reads that did not align. Transcript abundances were quantified by aligning these
rRNA-free files to a reference transcriptome (https://ftp.ncbi.nlm.nih.gov/genomes/
all/GCF/000/146/045/GCF_000146045.2_R64/GCF_000146045.2_R64_rna.fna.gz)
using Kallisto39. The resulting counts were rounded and TPMs recalculated based
on these rounded counts. Read count distributions for rounded count can be seen
in Supplementary Fig. 6a. Pairwise correlations based on the recalculated TPMs
produced high correlations between replicates (R > 0.99, Supplementary Fig. 6b).
Further analyses were based on these numbers and were performed using the R
programming language version 4.2.140 and the Tidyverse set of packages41. Dif-
ferential expression was performed using DESeq223 with the “apeglm”method of
normalization42. The GO term analysis was performed using Gene Ontology Slim
Term Mapper (https://www.yeastgenome.org/goSlimMapper). To determine if the
number of overlaps observed in Fig. 3c, d are unique, we generated an expected
value of overlaps by keeping the same number but randomizing the identity of
significant genes in each category and then recomputing the number of over-
lapping genes 10,000 times. The resulting distribution of overlaps +/−1 standard
deviation was less than the number of observed overlaps in both comparisons
(Fig. 3c and d). We used a chi-squared test to test for the independence of our
categories in each comparison. This is now presented in the Supplementary Fig. 6c.
circNC analysis. In total, 5 µg of yeast polyA RNA was first dephosphorylated
using Quick CIP (NEB) to eliminate potential 5′end phosphorylated RNA con-
tamination in the subsequent reactions. Typically, these reactions were performed
in a 100 µL volume with polyA-purified RNA dissolved in DEPC water, with 6 µL
of (CIP) in 10X rCutSmart™Buffer at 37 °C for 40 minutes. The dephosp horylated
RNAs were next purified using Monarch®RNA Cleanup Columns (NEB) as
described above and eluted in 30 µL of nuclease-free water. The dephosphorylated
RNAs are next divided into 3 equal fractions of 10 µL each and treated separately
with either no enzyme (Ctrl) or MDE (NEB) or Rai1. The MDE reaction was
performed using NEB’s protocol at 37 °C for 1 hour, whereas the Rai1 reaction was
performed in 50 µL volume with NEB buffer 3 and 0.5 µL RNAsin at 37 °C for 1 h.
The reaction products were next purified using Monarch®RNA Cleanup Columns
(NEB) and eluted in 15 µL DEPC water. All Ctrl, MDE, and Rai 1 treated RNAs
were next subjected to intramolecular ligation using T4 RNA ligase 1 (NEB).
Typically, the ligation reaction was performed in a 50 µL volume with 15 µL of Ctrl,
MDE, and Rai1 treated RNA following the manufacturer’s protocol in the presence
of ~25% PEG8000 at 25 °C for 16 hours. The reaction products were next purified
using Monarch®RNA Cleanup Columns (NEB) and eluted in 15 µL DEPC water
and reverse transcribed with M-MLV reverse transcriptase and gene-specific pri-
mers with gene name -P1 in Fig. 4a (sequence listed in Supplementary Table 3) as
per the manufacturer’s guidelines. PCR was performed with the primers (gene
name-P2 and gene name-P3 in Fig. 4a) listed in Supplementary Table 3 with Phire
Plant Direct PCR Master Mix (Thermo Scientific™). The PCR products were next
resolved onto either 1.5 or 2% Agarose-TAE gel, sliced, purified, and sequenced
using Amplicon-EZ (GENEWIZ (Azenta Life Sciences)) sequencing.
circNC sequencing analysis. Raw reads were processed for quality control and
adapter removal by fastp37. The analysis of this sequencing data was dependent on
our PCR products spanning the junction between the 5′and 3′ends of a transcript.
We chose primer locations near the start and end of the coding sequences for each
gene (Fig. 4a). However, because the UTR lengths for many yeast genes are
unknown and our read lengths are maximally ~500 bases (from 250 bp paired-end
reads), it was unclear if read 1 or 2 individually, or both would cover the junction.
To ensure that we were able to analyze the junction regardless of which read
covered it, we considered read 1 and read 2 separately, as well as the merged reads.
Reads were merged using BBmap’s bbmerge script43. Additionally, we expected
that if a read covered our junction, it should contain a polyA tail44. We used
cutadapt45 to filter our reads for those containing a stretch of at least 8 A’s in a row.
After filtering was complete, we counted the unique reads in each file (read 1, read
2, and merge) individually and used the top 5 most abundant unique reads for
multiple sequence alignments (using the R package msa46) which consisted of the
top 5 from an MDE sample and from a Rai1 sample. Manual, visual analysis of the
junction was carried out on the reads where the junction could be detected.
Sucrose gradient centrifugation. For polysome fractionation, exponentially
grown yeast cells in 500 mL YPD (OD
600nm
~2) were first treated with 100 µg/mL
cycloheximide for 10 minutes at room temperature. The cells were next harvested,
washed once in ice-cold 1x PBS, and lysed using mortar and pestle in liquid
nitrogen. The cell powder obtained after grinding was resuspended in polysome
buffer A (20-mM Tris-HCl pH7.4, 150-mM KCl, 3-mM MgCl2, 1-mM DTT, 0.5%
v/v NP-40 and EDTA- free complete protease inhibitor, Roche)47 and incubated at
4 °C for 30 min. Samples were then centrifuged for 10 min at 10,000 × gto remove
the cell debris. Approximately 40 A260 units were layered onto 10–50% w/v
sucrose gradients prepared in buffer A. The gradients were made with Gradient
Master 107 (Biocomp). These gradients were next centrifuged for 3 h at 39,000 rpm
and 4 °C in a Beckman-Coulter ultra-centrifuge using the SW40 rotor. After
centrifugation, fractions corresponding to each peak (Fig. 5a)–free pool, the 40 S,
60 S, 80 S, and polysomes were collected from the top of each gradient, by use of
Biocomp Gradient Station. The absorbance at 254 nm was measured during the
collection. Next, ribonucleoproteins from each fraction were precipitated using 1/
10th (v/v) 3 M sodium acetate (pH5.2) and 2X volume of 100% ethanol to get rid of
sucrose. The ribonucleoprotein pellet was next resuspended in 400 µL of DEPC
water and total RNA from each fraction was isolated using Trizol Reagent (Invi-
trogen) following manufacturers’instructions. For circNC analysis, polyA RNA
from each fraction was isolated as explained above.
Isolation of translationally active mitochondrial ribosomes. Translationally
active mitoribosomes were isolated as described before using C-terminally 3X
FLAG-tagged MRPS17 protein as a bait28. MRPS17-FLAG strain was generated
using homologous recombination using primers listed in Supplementary
Table3andusingplasmidpFA6a-6xGLY-3xFLAG-kanMX6asatemplate.5g
ofexponentiallygrownyeastcellpellet(OD600~2)inYPEG(1%yeastextract,
2% Peptone, 2% ethanol, and 2% Glycerol) was harvested and flash frozen in
liquid nitrogen. Cells were next lysed using a mortar and pestle in liquid
nitrogen. The cell powder obtained after grinding was resuspended in a lysis
buffer28,48 (10 mM Tris, pH 8.0, 50 mM NH4Cl, 10 mM MgCl
2
,0.5%CHAPS,
and 2X protease inhibitor cocktail (Complete, EDTA-free, Roche)) and incu-
batedat4°Cfor30min.Thelysatewasnextclarified by centrifugation for
10 min at 5000 × gto remove the cell debris. The clarified lysate was then
transferred to a fresh 15 mL centrifuge tube and the protein concertation was
assessed using a Bradford assay. 5 mg of protein lysate was added to the pre-
equilibrated (in 1X lysis buffer) anti-Flag M2 magnetic beads (Sigma). The
mixture was rotated end-over-end at 4 °C for 3 h. Next, the beads were washed
3X with wash buffer (10 mM Tris pH 8.0, 50 mM NH
4
Cl, 10 mM MgCl
2
,0.1%
CHAPS)for5minat4°CandtheFLAG-taggedproteinwaselutedbyincu-
bation with 200 μg/mL Flag peptide (Sigma). The eluate was next assessed for
thepresenceofMRPS17byWesternBlottingusinganti-FLAGantibody
(Sigma) (1;10,000). RNA from the eluate was isolated using Phenol-Chloroform
as described above.
Boronate affinity electrophoresis and Northern Blot analysis of in vivo NAD
capped transcripts after DNAzyme mediated RNA cleavage. Boronate affinity
electrophoresis was performed as described recently22.30μg of total cellular RNA
or the RNA retrieved from the mitoribosomes eluates (as described above) were
incubated with 1 μM of the corresponding DNAzyme in a 50 μL reaction con-
taining 10 mM Tris-HCl pH =8.0, 50 mM NaCl, 2 mM DTT, and 10 mM MgCl
2
.
Samples were denatured at 85 °C for 2 min and gradually cooled to 37 °C. MgCl
2
was added to a final concentration of 10 mM and incubated for 60 min at 37 °C.
Reactions were stopped with 100 μL of stop so lution (50 mM Tris pH 8.0, 20 mM
EDTA, and 0.1 µg/mL glycoge n), and RNA wa s precipitated with 500 μL ethanol
by incubating for 30 min at –80 °C followed by centrifugation for 30 min at
16,000 × gat 4 °C. The supernatant was removed, and the pellet was resuspended
in H
2
O.
NAD-capping was analyzed using DNAzyme-generated fragments of
mitochondrial RNAs. The cleaved RNA was resolved on 8% urea polyacrylamide
gels supplemented with 0.3% 3-acrylamidophenylboronic acid (Boron Molecular).
RNA was next transferred to a positively charged Nylon transfer membrane (GE
Healthcare Life Sciences) and incubated with a 32P-labeled DNA probe
complementary to the 5′-end fragments of target RNAs (Supplementary Table 3).
The probes were labeled using T4 polynucleotide kinase (NEB) and [γ-32P] ATP
(Perkin Elmer). Reaction products were visualized with Amersham Typhoon RGB
Biomolecular Imager (GE Healthcare Life Sciences).
Statistics and reproducibility. The present study utilized biological triplicates for
all samples in the NAD CapPro Seq and regular RNA seq experiments to ensure
statistical robustness and reproducibility. Statistical analysis was conducted using
Student’s t-test (single-tail), and chi-squared test to assess the independence of
categories in each comparison. The analyses were performed using the Tidyverse
package and other general statistics packages of the R programming language
version 4.2.1. The code used is available at https://github.com/shahlab/
NADcapPro. Differential expression analysis was conducted as described above
using DESeq2 with the “apeglm”as the method of normalization to control for
potential biases and improve the accuracy of the results.
ARTICLE COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-023-04774-6
12 COMMUNICATIONS BIOLOGY | (2023) 6:406 | https://doi.org/10.1038/s42003-023-04774-6 | www.nature.com/commsbio
Content courtesy of Springer Nature, terms of use apply. Rights reserved

Reporting summary. Further information on research design is available in the Nature
Portfolio Reporting Summary linked to this article.
Data availability
All unique materials and reagents generated in this study are available from the
corresponding author with a completed material transfer agreement. The sequencing
data is deposited at GSE217259. The code for analyses is available at https://github.com/
shahlab/NADcapPro. Any additional information required to reanalyze the data reported
in this paper is available from the corresponding author upon request. All data are
available in the main text or the supplementary materials.
Received: 19 December 2022; Accepted: 28 March 2023;
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Acknowledgements
We thank Xinfu Jiao and Haiyan Zheng for helpful discussions. The research was sup-
ported by NIH grant GM124976 to PS and NIH grant GM126488 to MK.
Author contributions
M.K., S.S. designed the experiments. S.S. and J.Y carried out the experiments and
computational analyses were carried out by J.F. and P.S. M.K. and S.S. wrote the
manuscript with input from all co-authors.
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Competing interests
P.S. is a Director at Ananke Therapeutics, a Scientific Advisory Board member of Trestle
Biosciences, and consults for Ribo-Therapeutics. All other authors declare no competing
interests.
Additional information
Supplementary information The online version contains supplementary material
available at https://doi.org/10.1038/s42003-023-04774-6.
Correspondence and requests for materials should be addressed to Sunny Sharma or
Megerditch Kiledjian.
Peer review information Communications Biology thanks Nan Liu, Sigitas Mikutis, and
the other, anonymous, reviewer(s) for their contribution to the peer review of this work.
Primary Handling Editor: Gene Chong.
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