Toll/interleukin-1 receptor (TIR) domain-containing proteins have NAD-RNA decapping activity

Abstract

The occurrence of NAD⁺ as a non-canonical RNA cap has been demonstrated in diverse organisms. TIR domain-containing proteins present in all kingdoms of life act in defense responses and can have NADase activity that hydrolyzes NAD⁺. Here, we show that TIR domain-containing proteins from several bacterial and one archaeal species can remove the NAM moiety from NAD-capped RNAs (NAD-RNAs). We demonstrate that the deNAMing activity of AbTir (from Acinetobacter baumannii) on NAD-RNA specifically produces a cyclic ADPR-RNA, which can be further decapped in vitro by known decapping enzymes. Heterologous expression of the wild-type but not a catalytic mutant AbTir in E. coli suppressed cell propagation and reduced the levels of NAD-RNAs from a subset of genes before cellular NAD⁺ levels are impacted. Collectively, the in vitro and in vivo analyses demonstrate that TIR domain-containing proteins can function as a deNAMing enzyme of NAD-RNAs, raising the possibility of TIR domain proteins acting in gene expression regulation.

Full text

Article https://doi.org/10.1038/s41467-024-46499-y
Toll/interleukin-1 receptor (TIR) domain-
containing proteins have NAD-RNA
decapping activity
Xufeng Wang
1,2,3,9
, Dongli Yu
4,5,6,9
, Jiancheng Yu
7
,HaoHu
1,2,3
,
Runlai Hang
1,2,3
, Zachary Amador
3
,QiChen
7,8
, Jijie Chai
4,5
&
Xuemei Chen
1,2
The occurrence of NAD+as a non-canonical RNA cap has been demonstrated in
diverse organisms. TIR domain-containing proteins present in all kingdoms of
life act in defense responses and can have NADase activity that hydrolyzes
NAD+. Here, we show that TIR domain-containing proteins from several bac-
terial and one archaeal species can remove the NAM moiety from NAD-capped
RNAs (NAD-RNAs). We demonstrate that the deNAMing activity of AbTir (from
Acinetobacter baumannii)onNAD-RNAspecifically produces a cyclic ADPR-
RNA, which can be further decapped in vitro by known decapping enzymes.
Heterologous expression of the wild-type but not a catalytic mutant AbTir in
E. coli suppressed cell propagation and reduced the levels of NAD-RNAs from a
subset of genes before cellular NAD+levels are impacted. Collectively, the
in vitro and in vivo analyses demonstrate that TIR domain-containing proteins
can function as a deNAMing enzyme of NAD-RNAs, raising the possibility of
TIR domain proteins acting in gene expression regulation.
Various chemical modifications decorate RNA biomolecules and
immensely expand the diversity of the transcriptome1,2.RNAmod-
ifications can occur internally or terminally3, and may alter the mole-
cular function, subcellular location, and stability of RNA4.Regardingthe
5′-terminal RNA structure, besides the most common triphosphophate
in prokaryotes and the N7-methylguanosine (m7G) cap in eukaryotes,
the redox cofactor nicotinamide adenine dinucleotide (NAD+)has
emerged as a non-canonical RNA cap structure in diverse organisms.
These include prokaryotes such as Escherichia coli (E. coli)5–7,Bacillus
subtilis (B. subtilis)8,Streptomyces venezuelae (S. venezuelae)5,and
Mycobacterium smegmatis9, and eukaryotes such as yeast10, mammalian
cells11,andplants
12–15. Recent studies have provided evidence that NAD-
capping of RNA also occurs in Archaea9,16, establishing the NAD cap
modification as ubiquitous in all three domains of life. Additionally,
other non-canonical RNA-caps, such as flavin adenine dinucleotide
(FAD), desphospho-coenzyme A (dpCoA), uridine diphosphate glucose
(UDP-Glc), uridine diphosphate N-acetyl glucosamine (UDP-GlcNAc),
and dinucleotide polyphosphate (Np
n
N), are also increasingly
described3,17–21.
The identity of NAD-capped RNAs (NAD-RNAs) varies in different
species. In bacteria, the protein-coding mRNAs are the major con-
stituents of the NAD-capped transcriptome, and somesmall regulatory
RNAs also tend to carry the NAD cap6–8,22. NAD-RNAs in archaea and the
eukaryotic kingdom also encompass various types of RNA species9,16,23,
underscoring the importance of studying the biogenesis and function
of NAD-RNAs.
Received: 6 October 2023
Accepted: 29 February 2024
Check for updates
1
State Key Laboratory for Protein and Plant Gene Research, Peking-Tsinghua Joint Center for Life Sciences, School of Life Sciences, Peking University, Beijing
100871, China.
2
Beijing Advanced Center of RNA Biology (BEACON), Peking University, Beijing 100871, China.
3
Department of Botany and Plant Sciences,
Institute of Integrative Genome Biology, University of California, Riverside, CA 92521, USA.
4
Institute of Biochemistry, University of Cologne, Cologne 50674,
Germany.
5
Max Planck Institute for Plant Breeding Research, Cologne 50829, Germany.
6
Dana-Farber Cancer Institute, Harvard Medical School, Boston, MA
02215, USA.
7
Department of Human Genetics, University of Utah School of Medicine, Salt Lake City, UT 84112, USA.
8
Molecular Medicine Program, Division of
Urology, Department of Surgery, University of Utah School of Medicine, Salt Lake City, UT 84112, USA.
9
These authors contributed equally: Xufeng Wang,
Dongli Yu. e-mail: xuemei.chen@pku.edu.cn
Nature Communications | (2024) 15:2261 1
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Thus far, two classes of NAD-RNA decapping enzymes have been
identified in prokaryotic or eukaryotic organisms24 (Supplementary
Fig. 1). Class-I decapping enzymes, such as the E. coli Nudix hydrolase
NudC, cleave the pyrophosphate bond within the NAD cap to liberate
the nicotinamide mononucleotide (NMN)6(Supplementary Fig. 1).
Another bacterial Nudix protein, BsRppH, catalyzes the decapping of
NAD-RNAs in Gram-positive B. subtilis by releasing NMN8. Homologs of
NudC in eukaryotic organisms, like Npy1 in yeast25 and Nudt12/16 in
mammalian cells26, have also been validated as the decapping
enzymes. Class-II decapping enzymes, such as the DXO/Rai1 family
enzymes in yeast, Arabidopsis, and mammals, remove the entire NAD
cap, producing monophosphorylated RNAs (p-RNAs)11,25,27,28 (Supple-
mentary Fig. 1). The 5′−3′exoribonucleases Xrn1 and Rat1 in the yeast
Saccharomyces cerevisiae also serve as NAD cap decapping (deNAD-
ding) enzymes, similar to DXO/Rai1,but primarily act onmitochondrial
NAD-RNAs29. Homologs of DXO/Rai1 have not been found in prokar-
yotic organisms. More recently, the human glycohydrolase CD38 has
been shown to convert NAD-RNAs into ADP-ribose-capped RNAs by
liberating nicotinamide (NAM) in vitro24 (Supplementary Fig. 1),
although its in vivo decapping activity still needs validation. Itis worth
noting that NudC and DXO/Rai1 can act on FAD-capped and dpCoA-
capped RNAs as well30–32 and are thus not specifictoNAD-RNAs.
NAD+can be catabolized by enzymes involved in cellular signaling
processes through cleaving the diphosphate bridge joining the two
nucleotides or the β-N-glycosidic bond linking ADP-ribose (ADPR) and
NAM33,34. Recent studies have shown that Toll/interleukin-1 receptor
(TIR) domain-containing proteins can act as NAD+-consuming enzymes
in bacteria34–37,mammals
38,39,andplants
38,40,41. They cleave free NAD+
molecules at the β-N-glycosidic bond into NAM and various forms of
ADPR isomers. This TIR domain-mediated NAD+hydrolase (NADase)
activity relies on a conserved catalytic glutamate (Glu/E) residue,
which is essential for the activation of downstream immune
responses34,36,38,40,41. However, it remains unexplored if TIR domain-
containing proteins have cleavage activity on the RNA NAD cap.
In this study, we provide evidence revealing that some TIR
domain-containing proteins remove the NAM moiety from NAD-RNAs.
This TIR domain-mediated deNAMing activity depends on the catalytic
Glu/E residue and is enhanced under conditions that promote the
oligomerization of TIR domain-containing proteins, such as in the
presence of molecularcrowding agents or when fused withthe tandem
Sterile Alpha Motif (tSAM) domain from the human SARM1 protein. We
identify the RNA product after AbTir (a TIR-domain containing protein
from Acinetobacter baumannii) treatment of NAD-RNA as a variant
form of cyclic ADPR-RNA (v-cADPR-RNA) that can be further decapped
by NudC or DXO/Rai1. TIR domain-containing proteins show decap-
ping activity specific to NAD-RNAs, distinct from other known decap-
ping enzymes. We further demonstrate that AbTir is functional in
bacteria –its inducible expression in E. coli suppresses growth and
significantly decreases the levels of both free NAD+and NAD-RNAs.
SPAAC-NAD-Seq profiling indicates that NAD-RNAs produced from a
small subset of genes involved in “molecule transport process”and
“oxidoreductase activity”are targeted by AbTir in E. coli,furthercon-
firming the in vivo deNAMing activity of TIR domains-containing
proteins. Finally, we find that an archaeal TIR domain-containing pro-
tein, TcpA, also exhibits deNAMing activity on NAD-RNA, extending
the deNAMing activity of TIR domains to archaea. Our findings reveal
NAD-RNA deNAMing as a m olecular function of TIR domain-containing
proteins and implicate a role of TIR domain proteins in gene expres-
sion regulation.
Results
Prokaryotic TIR domain-containing proteins show deNAMing
activity on NAD-RNAs
The NADase activity of TIR domain proteins prompted us to test
whether they also act on NAD-RNAs. We first expressed a suite of
bacterial TIR domain-containing proteins asrecombinant proteins in E.
coli, including AbTir, BtTir, BtpA, TcpC, and TcpF from pathogenic
bacteria and TirS and PdTir from non-pathogenic bacteria (Fig. 1a;
Supplementary Fig. 2a). All tested TIR domain-containing proteins
hydrolyzed free NAD+to varying degrees in vivo and in vitro (Supple-
mentaryFig. 2b, c), showing NADase activity as reported35. Next, to test
whether these TIR domain-containing proteins can hydrolyze the RNA
NAD cap, we synthesized a 32-nucleotide (nt) NAD-RNA by in vitro
transcription using T7 RNA polymerase (Supplementary Fig. 3) and
incubated it with the purified TIR domain-containing proteins. The
reaction products were subsequently separated by acryloylamino-
phenyl boronic acid polyacrylamide gel electrophoresis (APB-PAGE),
which can selectively retard the migration of capped RNAs by reacting
with the vicinal diol of the cap nucleoside42. The results revealed that
AbTir from Acinetobacter baumannii, BtpA from Brucella abortus,and
PdTir from Paracoccus denitrificans, but not the other tested bacterial
TIR domain-containing proteins, showed decapping activity on the
NAD-RNA under the assay conditions employed (Fig. 1b). Mutating the
putative catalytic Glu/E residues from the TIR domain-containing
proteins abolished the activity on both free NAD+(i.e., NADase activity)
and the NAD-RNA (i.e., decapping activity) (Fig. 1c–e).
Time-course assays with AbTir further substantiated the enzy-
matic decapping activity and showed that AbTir cleaved free NAD+
slightly faster than NAD-RNA in vitro (Fig. 2a, b; Supplementary Fig. 4).
As expected, NAD+depletion accompanying NAM accumulation was
detected by high-performance liquid chromatography coupled with
mass spectrometry (HPLC-MS) after incubating AbTir with free NAD+
(Fig. 2c). Consistently, NAM was released when AbTir was incubated
with an in vitro transcribed NAD-RNA (Fig. 2c), suggesting that AbTir
cleaved the β-N-glycosidic bond within the NAD cap to release the
NAM moiety (this activity is referred to as deNAMing hereafter;
Fig. 2d). NAD-capQ is another method for the quantification of NAD-
RNAs among total RNAs by measuring the amount of NAD+released
from RNA after nuclease P1 digestion43. To validate the decapping
ability of AbTir on native NAD-RNAs, we subjected cellular total RNAs
from either Arabidopsis or E. coli to AbTir treatment followed by
nuclease P1 digestion and NAD-capQ measurement (Fig. 2e). As com-
pared to the catalytically inactive AbTir-E/A, wild-type AbTir caused a
significant reduction in the levels of the NAD cap from both in vitro
transcribed and native NAD-RNAs (Fig. 2f). Taken together, these
results revealed that some bacterial TIR domain-containing proteins
can process NAD-RNAs by removing the NAM moiety from the NAD
cap, showing a deNAMing activity on in vitro transcribed and native
NAD-RNAs.
Oligomerization enhances the deNAMing activity of AbTir
Next, we evaluated the parameters of the deNAMing assay to optimize
the deNAMing reaction. Oligomerization of TIR domains is funda-
mental for the NADase activity38,41,44–47.Wefirst examined the
deNAMing activity of AbTir with different concentrations of the pur-
ified protein and observed that the deNAMing activity was non-linearly
increased with protein concentrations (Supplementary Fig. 5a, b). This
observation implied that self-association might alsobe required forthe
NAD-RNA deNAMing activity of AbTir, consistent with prior findings
that oligomerization of TIR domain proteins enhances the NADase
activity on NAD+47–49. Polyethylene glycol (PEG), as a molecular
crowding agent, can simulate a crowded environment inside cells, and
promote the self-association of TIR domains38,41.Wethusapplieddif-
ferent types of PEGs in the reaction and found that the deNAMing
activity was improved in the presence of certain PEGs (Supplementary
Fig. 5c). The same amount of NAD-RNAs was cleaved by AbTir in 2 h in
the presence of PEG6000 as that in 16h in the absence of PEG6000
(Supplementary Fig. 5d). The tandem Sterile Alpha Motif (tSAM)
domain of human SARM1 (Sterile alpha and HEAT/Armadillo motif)
enhances the NADase activity of TIR domains by facilitating their
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 2
Content courtesy of Springer Nature, terms of use apply. Rights reserved

multimerization39,50,51. We fused this tSAM domain (hSARM1tSAM;resi-
dues 409-561aa) to AbTirTIR (TIR domain of AbTir, residues 133-266aa)
(Supplementary Fig. 5e) and purified the fusion protein hSARM1tSAM-
AbTirTIR from E. coli (Supplementary Fig. 5f). As expected, fusion of
hSARM1tSAM to AbTirTIR significantly enhanced the deNAMing activity of
AbTir (Supplementary Fig. 5g). Collectively, these results indicated
that self-association of AbTir plays a pivotal role in bolstering its
deNAMing activity on NAD-RNAs.
Identification of the RNA product of deNAMing by AbTir
TheRNAproductafterAbTircleavageshowedfastermigrationon
APB-PAGE than in vitro transcribed ADPR-RNA (Fig. 1c), which
prompted us to investigate the identity of this cleavage product. As a
NADase enzyme, AbTir cleaves free NAD+into NAM and a cADPR iso-
mer known as v-cADPR or 2′-cADPR, meaning that AbTir has a cyclase
activity in addition to the cleavage activity35,37,47. Thus, we assume that
the RNA product of AbTir might possess a cyclic ADPR cap (Fig. 3a). A
recent study demonstrated that a conserved tryptophan of AbTir is
required for the cyclization of ADPR after the hydrolysis reaction with
free NAD+47. Mutating this residue from tryptophan (Trp/W) to alanine
(Ala/A) significantly reduced, but did not abolish, the production of 2′-
cADPR, compared to wild-type AbTir47. We thus hypothesized that
mutating the tryptophan residue will result in the generation of non-
cyclic ADPR-RNA. We generated the AbTir-W204A mutant and per-
formed the NADase and decapping assays with the purified protein
(Fig. 3b, c). Consistent with the previous report47, a significant reduc-
tion in the NADase activity was observed (Fig. 3b). The decapping
activitywasalsoreduced,asreflected by the input NAD-RNA being
incompletely consumed by AbTir-W204A (Fig. 3c). Interestingly, two
RNA products were produced by AbTir-W204A, with one showing the
same migration as the AbTir product and the other showing the same
migration as the in vitro transcribed ADPR-RNA (Fig. 3c), suggesting
that this tryptophan residue plays an important role in determining the
nature of the AbTir NAD-RNA deNAMing product. This observation
also implied that the RNA product from AbTir treatment should be a
certain type of cyclic ADPR-RNA. Unfortunately, we failed to obtain
cADPR-RNA by in vitro transcription using cADPR in place of ATP and
v-cADPR is not commercially available for use in in vitro transcription,
ab
c
de
absorbance (450 nm)
elution buffer
****
absorbance (450 nm)
elution buffer
****
PdTir
PdTir-E/A
BtpA
BtpA-E/A
0
0.5
1.0
1.5
2.0
2.5
3.0
0
0.5
1.0
1.5
2.0
2.5
3.0
absorbance (450 nm)
elution buffer
AbTir
AbTir-E/A
AbTir
AbTir-E/A
NAD-RNA ADPR-RN
A
****
0
0.5
1.0
1.5
2.0
2.5
3.0
cleavage product
2141 275
TirS (S. aureus)
N
133 266
CAbTir (A. baumannii)
169 303
TcpC (E. coli)
142 275
BtpA (B. abortus)
164 297
PdTir (P. dentrificans)
140
TcpF (E. faecalis)
TIRCC
TIRCC
TIR
TIR
TIR
TIR
pathogenic bacteria
non-pathogenic bacteria
287
BtTir (B. thetaiotaomicron)TIR
31 99
156
3
94
AbTir
BtTir
TcpC
TIR
BtpA
TcpF
PdTir
TirS
NAD-RNA
ppp-RNA
cleavage product
NAD-RNA ADPR-RNA
ppp-RNA
cleavage product
NAD-RNA ADPR-RNA
ppp-RNA
cleavage product
Boronate gel
Boronate gel
Boronate gel Boronate gel
NAD-RNA
+++++
protein
-
+++
NAD-RNA
-++ -
+
ppp-RNA
+----
ADPR-RNA
---+-
protein
-- -
PdTir
PdTir-E/A
NAD-RNA
++ -
+
ADPR-RNA
-- +-
protein
--
BtpA
BtpA-E/A
NAD-RNA
++ -
+
ADPR-RNA
-- +
-
protein
--
Fig. 1 | Bacterial TIR domain-containing proteins cleave the NAD cap of NAD-
RNAs. a Diagrams of the tested bacterial TIR domain-containing proteins. The
numbers above indicate the start and end amino acids of the annotated domains.
CC, coiled coil; TIR, Toll/interleukin-1 receptor. bAn acryloylaminophenyl boronic
acid (APB) gel showing the outcome of decapping assays, in whichvarious bacterial
TIR domain-containing proteins were incubated with an in vitro transcribed NAD-
RNA. Three TIR domain-containing proteins (AbTir, BtpA, and PdTir) showed
activities toward the NAD-RNA. A reaction without purified protein added was
included for comparison. The positions of NAD-RNA, ppp-RNA and the cleavage
productare indicated.Assays for the enzymatic activities of AbTir(c), BtpA (d), and
PdTir (e) and their corresponding catalytic mutants. Both the NADase activities on
free NAD+(left) and the decapping activities on the NAD-RNA (right) were depen-
dent on the putative catalytic Glu/E residues. The relative NAD+content after the
NADase assay was measured with the NAD/NADH Quantitation Kit by monitoring
theabsorbancevaluesat450nm.Horizontalbarsrepresentmedianvaluesoffive
independent experiments (n=5);[**] P≤0.01 (calculated by the non-parametric
Mann-Whitney U-test, two-sided). In vitro transcribed ppp-RNA and ADPR-RNA
were used asmarkers for comparison to the NAD-RNA cleavage products. Exact P-
values and source data are provided as a Source Data file.
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 3
Content courtesy of Springer Nature, terms of use apply. Rights reserved

which impeded the ability to narrow down the identity of the RNA
product by APB-PAGE.
It is known that adenosine diphosphate-ribosyl cyclase (ADPRC)
purified from Aplysia californica is a bifunctional enzyme that releases
NAM from NAD+and cyclizes ADPR to cADPR52.Anotherbifunctional
enzyme, human CD38, catalyzes the hydrolysis of NAD+as a glycohy-
drolase, but the cyclase activity is weak, resulting in 97% of the pro-
ducts being ADPR53–55. Both ADPRC and CD38 are able to remove the
NAM moiety from in vitro transcribed NAD-RNA6,24. The RNA product
of ADPRC-catalyzed NAD-RNA deNAMing was not determined. While
Abele et al. reported ADPR-RNA as one of the cleavage products of
CD38 on in vitro transcribed NAD-RNA24, they did not explore other
potential products, such as cADPR-RNA and v-cADPR-RNA. We ana-
lyzed the reaction products of AbTir, ADPRC, and CD38 on free NAD+
by HPLC-MS, and found that v-cADPR47,56 and ADPR were detected for
AbTir and CD38, respectively (Supplementary Fig. 6), as
reported47,53,54,56. However, a clear peak corresponding to the ADPR
standard was detected for ADPRC (Supplementary Fig. 6), not cADPR
as previously reported52. After knowing the products produced by
these enzymes on free NAD+, we compared the products of AbTir,
ADPRC and CD38 on NAD-RNA. In APB-PAGE, the RNA products of
ADPRC and CD38 showed the same migration as the in vitro tran-
scribed ADPR-RNA (Fig. 3d). To unambiguously determine the iden-
tities of the RNA products after ADPRC and CD38 cleavage, we
established an HPLC-MS analysis pipeline. The in vitro transcribed
NAD-RNA was first incubated with either ADPRC or CD38 to release the
NAM moiety from the NAD-RNA, and then the RNA products were
further digested by nuclease P1 into single nucleotides, which were
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10.0 11.0 12.0 13.0 14. 0 15.0 16.0 17.0 18.0 19.0
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
(x1,000,000)
1.0 2.0
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10.0 11.0 12.0 13.0 14.0 15.0 16.0 17.0 18.0 19.0
0.0
1.0
2.0
3.0
4.0
5.0
6.0
7.0
8.0
9.0
(x10,000)
c
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9. 0 10.0 11.0 12.0 13.0 14.0 15.0 16.0 17.0 18.0 19.0
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
2.00
2.25
2.50
2.75
(x10,000)
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10.0 11. 0 12.0 13.0 14.0 15.0 16.0 17.0 18.0 19.0
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
2.00
2.25
2.50
2.75
(x10,000)
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10.0 11.0 12.0 13. 0 14.0 15.0 16.0 17.0 18.0 19.0
0.0
1.0
2.0
3.0
4.0
5.0
6.0
7.0
8.0
9.0
(x10,000)
0
0.5
1.0
1.5
2.0
0
0
0,0
00
,1x
0
2.0
4.0
6.0
8.0
000,0
1x
0
2.0
4.0
6.0
8.0
000,0
1x
0
0.5
1.0
1.5
2.0
000,0
1x
0
0.5
1.0
1.5
2.0
0
00
,0
1x
NAD+
2143657
retention time (min)
intensity
a
d
AbTir
AbTir-E/A
0246810121416
NAD-RNA
ppp-RNA
time (hour)
cleavage product
NAD-RNA
ppp-RNA
0
20
40
60
80
100
120
140
02468101214
100μM NAD++AbTir
NAD-RNA + AbTir NAD-RNA + AbTir-E/A
remaining NAD+or NAD-RNA (%)
100μM NAD++ AbTir-E/A
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10.0 11.0 12.0 13.0 14. 0 15.0 16.0 17.0 18.0 19.0
0.0
1.0
2.0
3.0
4.0
5.0
6.0
(x100,000)
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10.0 11.0 12. 0 13.0 14.0 15.0 16.0 17.0 18.0 19.0
0.0
1.0
2.0
3.0
4.0
5.0
6.0
(x100,000)
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10. 0 11.0 12.0 13.0 14.0 15.0 16.0 17.0 18.0 19.0
0.0
0.5
1.0
1.5
2.0
2.5
3.0
(x100,000)
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10.0 11.0 12.0 13.0 14.0 15.0 16. 0 17.0 18.0 19.0
0.0
0.5
1.0
1.5
2.0
2.5
3.0
(x100,000)
1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10. 0 11.0 12.0 13.0 14.0 15.0 16.0 17.0 18.0 19.0
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
2.00
(x10,000,000)
0
0.5
1.0
1.5
2.0
0
1.0
2.0
3.0
4.0
0
1.0
2.0
3.0
4.0
0
2.0
4.0
6.0
8.0
0
2.0
4.0
6.0
8.0
buffer + NAD+
AbTir + NAD+
NAD-RNA + P1
AbTir + NAD-RNA + P1
b
16 8109
NAM
absorbance (450nm)
0
0.5
1.0
1.5
2.0
*
AbTir+P1
AbTir-E/A+P1
0
0.5
1.0
1.5
**
AbTir+P1
AbTir-E/A+P1
0
0.1
0.2
0.3
0.4
AbTir+P1
AbTir-E/A+P1
**
NAD-RNA Arabidopsis
total RNA
E. coli
total RNA
OH
O
O
O
O
O
P
P
O
+
RNA
NAD-RNA
NAD+
measurement
NAM AbTir
Nuclease P1
OH OH
N
NH2
O -
O
O -
NN
N
N
NH2cleavage product
f
standard
absorbance (450nm)
absorbance (450nm)
retention time (min)
incubation time (hour)
e
0214365781090
Fig. 2 | AbTir exhibitsdeNAMing activity on NAD-RNAs. a AtimecourseofNAD-
RNA decapping by wild-type AbTir and its catalytic mutant AbTir-E/A. An in vitro
transcribed NAD-RNA was incubatedwith AbTir (top panel) or its catalytic mutant
AbTir-E/A (bottom panel) forthe indicated duration and RNAsin the reaction were
resolved in denaturing APB gels. bComparison of the reaction kinetics of the
NADase activity on free NAD+and the decapping acticity on NAD-RNA by AbTir.
The catalytic mutant AbTir-E/A was included as a negative control. For each time
point, the remaining substrates were analyzed from three independent experi-
ments with error bars representing mean ± SD (n= 3, Supplementary Fig. 4).
cHPLC-MS analysis demonstrating that free NAD+or the NAD cap from the NAD-
RNA was largely destroyed, accompanied by nicotinamide (NAM) accumulation
after AbTir treatment. For the detection of NAD+in RNA, theRNA was treated with
nuclease P1 (P1) before HPLC-MS analysis. The three product ions from NAD+
([M + H]+664.00 > 136.00, 664.00 > 427.90, 664.00 > 523.95) and NAM ([M + H]+
123.00 > 80.00, 123.00 > 78.00, 123.00 > 53.00) are indicated by black, pink and
blue lines, respectively. dDiagram of NAD-RNAand its deNAMing cleavage site by
AbTir. The cleavage site in the NAD cap is marked with a blue dashedline, and the
NAM structure is highlighted in red color. eThe experimental pipeline showing
NAD cap breakdown from NAD-RNA by pretreatment with AbTir. After AbTir
cleavage, the NAD cap should be destroyed and should not be detected after
nuclease P1 digestion of the RNA. fMeasurement of NAD cap content using the
pipeline in (e) after AbTir (orange) or AbTir-E/A (light blue) treatment. Anin vitro
transcribed NAD-RNA (0.5 μg) was used as a positive control. 500 μg total RNA
from Arabidopsis or E. coli was used to validate the deNAMing activity of AbTir.
Error bars represent mean ± SD, which were calculated from three biologically
independent samples (n= 3); [**] P≤0.01; [*] P≤0.05 (calculated by the Student’s
t-test, two-sided). Exact P-values and source data are provided as a Source
Data file.
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 4
Content courtesy of Springer Nature, terms of use apply. Rights reserved

subjected to HPLC-MS analysis (Fig. 3e).Thisanalysisdetectedboth
ADPR and cADPR for reactions with CD38 and ADPRC (Fig. 3f),
meaning that both ADPR-RNA and cADPR-RNA are the RNA products
upon CD38 or ADPRC treatment. With this pipeline (Fig. 3e), we found
that the same AbTir NADase product, i.e., v-cADPR, was detected in the
AbTir deNAMing reaction on the NAD-RNA (Fig. 3f; Supplementary
Fig. 6), indicating that v-cADPR-RNA was the specificRNAproductof
AbTir-mediated NAD-RNA deNAMing.
AbTir specifically cleaves the RNA NAD cap in vitro as an initial
step for decapping
Various moieties, such as the triphosphate group, NAD+, dpCoA, FAD,
and Np
n
N, have been reported to be covalently attached to the 5′
terminus of RNA and might affect RNA stability3,17–19. In vitro and
in vivo, NAD-capped RNAs can be decapped by Nudix and DXO/Rai1
family enzymes, originally identified in disparate prokaryotic and
eukaryotic organisms6,11,24. Both classes of enzymes decap NAD-RNAs
but also decap RNAs with other non-canonical caps, such as FAD and
dpCoA30,31,57,58. The primary product of decapping isp-RNA, which can
undergo rapid RNA decay.
Toexaminethesubstratespecificity of AbTir, we used a set of
in vitro transcribed RNAs with non-canonical caps (Supplementary
Fig. 3). We found that NudC and DXO can indiscriminately cleave the
caps of NAD-RNA, FAD-RNA, dpCoA-RNA, and ADPR-RNA (Fig. 4a).
NudC also showed robust cleavage activity on Ap
4
A-RNA, while DXO
did not. Among all the substrate RNAs tested, AbTir only acted on
NAD-RNA (Fig. 4a).
Because in vitro transcribed ADPR-RNA can be decapped by NudC
andDXO,wetestedwhetherthev-cADPR-RNA generated by AbTir can
be further decapped by NudC or DXO. We added NudC and DXO
directly to the reaction after the NAD-RNA was incubated with AbTir.
We observed that the v-cADPR-RNA was further cleaved into p-RNA by
NudC or DXO (Fig. 4b). Therefore, TIR domain-containing proteins can
specifically remove NAM from the NAD cap, and a second decapping
step is required to produce p-RNA (Fig. 4c), suggesting that these
enzymes might work together to specifically decap NAD-RNAs. A
0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
2.00
(x100,000)
0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
2.00
(x100,000)
0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
(x10,000,000)
NAD-RNA
ADPR, cADPR, v-cADPR ...
NAM
AbTir/
ADPRC/
CD38
nuclease
P1
RNA products
(ADPR-RNA, cADPR-RNA, v-cADPR-RNA ...)
d
e
f
abc
0
1
2
3
4
5
absorbance (450nm)
****
*
AbTir
elution buffer
AbTir-W204A
OH
O
O
O
O
O
P
P
OH OH
O -
O
O -
NN
N
N
NH2
O
OH
OH
O
O
O
O
O
P
P
OH OH
O -
O
O -
NN
N
N
NH2
O
ADPR-RNA cADPR-RNA v-cADPR-RNA
O
O
O
O
O
O
P
P
OH OH
O -
O
O -
NN
N
NH2
O
N
0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
(x10,000,000)
0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0
0.0
0.5
1.0
1.5
2.0
2.5
3.0
3.5
4.0
4.5
(x1,000,000)
0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5 10.0
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
(x10,000,00 0)
0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
2.00
2.25
2.50
(x10,000,000)
0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5
0.00
0.25
0.50
0.75
1.00
1.25
1.50
1.75
2.00
(x100,000)
ADPR cADPR
retention time (min)retention time (min)
intensity intensity
intensity
intensity
0
5.0
10.0
15.0
20.0
0
2.5
5.0
7.5
10.0
123456
789123456789
0
5.0
10.0
15.0
20.0
0
2.5
5.0
7.5
10.0
0
5.0
10.0
15.0
20.0
0
5.0
10.0
15.0
20.0
0
5.0
10.0
15.0
20.0
0
2.5
5.0
7.5
10.0
0
00,0
00,1
x
00
0,0
00,1
x
0
0
0
,0
00,1
x
0
00,0
00,1x
standard
CD38
ADPRC
AbTir
NAD-RNA
cleavage product 1
cleavage product 2
ppp-RNA
Boronate gel
Boronate gel
AbTir
AbTir-W204A
NAD-RNA
++ -+
ADPR-RNA
-- +
-
protein
--
ADPR-RN
A
v-cADPR
AbTir-E/A
AbTir
NAD-RNA
++++
ADPR-RNA
-
--
protein
-
CD38
-
+
+
-
-
ADPRC
NAD-RNA
cleavage product 1
cleavage product 2
ppp-RNA
ADPR-RNA
-
0
0
0,
0
0
0
,
1
x
00
0,
0
1x
000,
01
x
00
0
,
0
1x
Fig. 3 | Identification of the RNA product of NAD-RNA deNAMing by AbTir.
aDiagrams showing the possible products (ADPR-RNA, cADPR-RNA, and v-cADPR-
RNA) of NAD-RNA deNAMing by TIR domain-containing proteins. bNADase assays
(see Methods) to measure the hydrolytic activity ofwild-type AbTir and its W204A
mutant on free NAD+. The W204A mutation significantly reduced the NADase
activity. Error bars represent mean± SD, which was calculated from three inde-
pendent experiments (n= 3); [**] P≤0.01 (calculated by the Student’st-test, two-
sided). cNAD-RNA decapping assays for wild-type AbTir and its W204A mutant.
The AbTir-W204A mutant showedreduced deNAMingactivity compared with wild-
type AbTir but generated a new cleavage product corresponding to the in vitro
transcri bed ADPR-RNA. dAn APB gel showing theproducts of NAD-RNAdecapping
by AbTir, ADPRC and CD38. An in vitrotranscribed NAD-RNA was used as the
substrate. ADPRC and CD38 converted the NAD-RNA to a product of the same
mobility as the in vitro transcribed ADPR-RNA. The RNA product from AbTir had
higher mobility in the APB gel than the products from ADPRC or CD38, or the
in vitro transcribed ADPR-RNA. eA pipeline used for identifying the deNAMing
products by AbTir, ADPRC, and CD38. fHPLC-MS analyses following the pipeline in
(e) demonstrated that both ADPR-RNA and cADPR-RNA were the products gener-
ated by ADPRC and CD38, whilev-cADPR-RNA wasproduced by AbTir.Commercial
ADPR and cADPR were used as standards for the HPLC-MS analysis. The three
product ions of cADPR ([M + H]+541.80 > 136.15, 541.80> 427.90, 541.80 > 347.90)
and ADPR ([M+ H]+559.80 > 136.05,559.80 > 347.95, 559.80> 427.95) are indicated
by black, pink and blue lines, respectively. Exact P-values and source data are
provided as a Source Data file.
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 5
Content courtesy of Springer Nature, terms of use apply. Rights reserved

recent report showed that ADPR-RNA exists in archaea and can be
generated from NAD-RNA through the spontaneous loss of NAM16.Our
findings suggest that TIR-catalyzed NAM removal is another mechan-
ism to generate RNAs capped by ADPR or its variants (Fig. 4c).
We compared the rate of NAD-RNA decapping by these three
enzymes through a time-course assay. The results showed that AbTir
could hydrolyze the NAD cap at a similar rate as NudC and DXO
(Fig. 4d; Supplementary Fig. 7). These results suggest that multiple
mechanisms might exist forNAD-RNA decapping in vivo.In addition to
Nudix hydrolases and DXO/Rai1 family enzymes, TIR domain-
containing proteins may also play a role in removing the NAD cap.
AbTir deNAMs native NAD-RNAs
Having shown thatAbTir removes NAM from in vitro transcribed NAD-
RNAs, we sought to determine whether AbTir deNAMs cellular NAD-
RNAs. In previous studies, it was shown that NAD-RNAs can be selec-
tively biotinylated by Copper-catalyzed azide-alkyne cycloaddition or
strain-promoted azide-alkyne cycloaddition (SPAAC) click chemistry
and biotinylated RNAs can be detected by gel blots or captured for
high throughput sequencing6,7,15.UnlikeNAD-RNA,wefoundthat
ADPR-RNA was not biotinylated by the SPAAC reaction (Supplemen-
tary Fig. 8). Based on this observation, we envisioned that NAD-RNAs
could not be biotinylated if pre-treatedwith AbTir (Fig. 5a). To test this,
we treated an in vitro transcribed NAD-RNA and native mRNAs isolated
from Arabidopsis and E. coli with wild-type AbTir or catalytically
inactive AbTir-E/A, and then performed the SPAAC reaction followed
by gel analysis. Treatment with wild-type AbTir significantly reduced
the detection of biotin-linked RNAs, while treatment with AbTir-E/A
had no effect on signal detection (Fig. 5b, c). This showed that AbTir
can deNAM Arabidopsis or E. coli NAD-RNAs in vitro.
To identify the NAD-RNA species acted upon by AbTir, we per-
formed SPAAC-NAD-Seq15 using E. coli ribosomal RNA (rRNA)-depleted
RNA after AbTir or AbTir-E/A treatment. The results of SPAAC-NAD-Seq
were highly reproducible among biological replicates (Supplementary
Fig. 9). After AbTir treatment, we identified only 12 NAD-RNA-
producing genes with E. coli rRNA-depleted RNA, whereas after treat-
ment with AbTir-E/A, we identified 1,057 NAD-RNA-producing genes
(Ratio of NAD-RNA-Seq/regular RNA-Seq ≥2&FDR≤0.05; Fig. 5d;
Supplementary Data 1 and 2), suggesting that almost all NAD-RNAs are
sensitive to deNAMing activity by AbTir in vitro.
In a previous report using NAD tagseq II, a total of 279 NAD-RNA-
producing genes were identified in E. coli7. Among these genes, 248
had expressed homologs in the BL21 (DE3) strain used in our study,
and 165 (~67%) of them were also identified as NAD-RNA-producing
genes in our SPAAC-NAD-Seq analysis (Supplementary Fig. 10a). This
demonstrates the accuracy and robustness of our SPAAC-NAD-Seq
profiling. The increased number of NAD-RNA-producing genes iden-
tified in our analysis may be attributed to the limited sensitivity of
Oxford nanopore sequencing used in NAD tagSeq II, as well as differ-
ential expression of cognate genes under different conditions. The
newly identified 1,057 NAD-RNA-producing genes are predominantly
protein-coding genes, with a few from non-coding RNAs (Supple-
mentary Fig. 10b). These NAD-RNA-producing genes tend to be shorter
and have lower expression levels (Supplementary Fig. 10c, d). Gene
ontology (GO) analysis revealed enrichment in functional categories
related to “RNA metabolic process”and “regulation of gene expres-
sion”(Supplementary Fig. 10e). We specifically focused on the top 20
NAD-RNAs identified in the NAD tagSeq II assay and found that 15 (75%)
of them were highly enriched as NAD-RNAs after AbTir-E/A treatment
(Fig. 5f). Among the remaining 5 NAD-RNAs, four of them (sibD,amyA,
DXO
NudC
AbTir-E/A
AbTir
NudC
AbTir
NAD-RNA
+++
+-
+
+--
incubate first
DXO
AbTir
NAD-RNA
+++
+-
+
+--
incubate first
a
b
c
NAD-
v-cADPR-
cADPR-
ADPR-
or
or
p-
TIR protein
(eg. AbTir)
Nudix
exonuclease
NAM
NMN
DXO/Rai1
d
010306090120150
-50
0
50
100
150
incubation time (min)
DXO
NudC
AbTir function?
NAD
remaining NAD-RNA (%)
ppp-RNA
ppp-RNA
NAD-RNA
+++++
DXO
NudC
AbTir-E/A
AbTir
FAD-RNA
DXO
NudC
AbTir-E/A
AbTir
dpCoA-RNA
Ap
4
A-RNA ADPR-RNA ppp-RNA
NAD-RNA
p/ppp-RNA
v-cADPR-RNA FAD-RNA
ppp-RNA
dpCoA-RNA
p/ppp-RNA
ADPR-RNA
p/ppp-RNA ppp-RNA
Ap
4
A-RNA
p/ppp-RNA
protein
-
protein
-
protein
-
++++++++++
+++++++++++++++
NAD-RNA
v-cADPR-RNA
p/ppp-RNA
NAD-RNA
v-cADPR-RNA
p/ppp-RNA
Fig. 4 | Specificity of AbTir on non-canonical RNA caps in comparison to NudC
and DXO. a APBgels showing in vitrotranscribed RNAs with various non-canonical
caps (NAD, FAD, dpCoA, Ap
4
A, ADPR,and triphosphate) and decapping activitiesof
AbTir, NudC, and DXO on these RNAs. Reactions without the enzymes were
included for comparison. The red arrows represent the positions of the indicated
RNA species in APB gels. bAPB gels showing the combined effects of AbTir and
NudC/DXOenzymes. ppp-RNA was loaded as a marker to show the position of the
final cleavage product. NAD-RNA was incubated first with AbTir for the deNAMing
assay, and then NudC or DXO was added into the reaction for further incubation.
cSchematic of NAD-RNA hydrolysis by Nudix family enzymes, DXO/Rai1 deNAD-
ding enzymes, and TIR-domain deNAMing enzymes. dRates of NAD-RNA decap-
ping by AbTir, NudC, and DXO. Data points represent mean ± SD. For each time
point, the reaction kinetics was analyzed from three independent decapping assays
(Supplementary Fig. 7). The y-axis indicates the proportions of remaining NAD-
RNAs after decapping reactions. In each reaction, 500ng NAD-RNA was incubated
with 1 0 μM AbTir/NudC/DXO. Source data are provided as a Source Data file.
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 6
Content courtesy of Springer Nature, terms of use apply. Rights reserved

ykgS,andyfjI) did not have cognate genes in the E. coli BL21 (DE3) strain
and one of them (yfcO) was not detected as an NAD-RNA. However,
after treatment with AbTir, the enrichment of these NAD-RNAs was
significantly reduced (Fig. 5f; Supplementary Fig. 11). This indicates the
strong decapping activity of AbTir on native RNA.
AbTirtargetsasubsetofNAD-RNAsinE. coli
To determine the in vivo decapping activity of AbTir, we expressed the
wild-type AbTiror the catalytically inactive AbTir-E/A (Fig. 6a) in the E.
coli BL21 (DE3) strain. We monitored cell proliferation by measuring
the OD values at 600 nm (OD600) after AbTir induction with different
concentrations of IPTG. E. coli cells expressing AbTir showed
decreased cell proliferation compared to those expressing AbTir-E/A
(Fig. 6b; Supplementary Fig. 12a–c). This decrease in cell proliferation
may be due to the consumption of NAD+in E. coli cells by AbTir, as
previously reported35,47 and confirmed in our measurements (Fig. 6c;
Supplementary Fig. 12d). We sought to determine whether AbTir also
caused NAD-RNA deNAMing in bacteria. To this end, we first measured
the endogenous NAD+content at different time points after IPTG
addition (Fig. 6d; Supplementary Fig. 13), and found a time point, at
which AbTir accumulated but levels of free NAD+were not affected
(Fig. 6d, e). At this time point (15 min after induction with 0.1 mM
H2O
AbTir
AbTir-E/A
+
-
-
+
-
-+
-
-
+
-
-
+
-
-+
-
-
(1) (2)
b
c
+
-
-
+
-
-+
-
-
(3)
NAD-RNA
or
native mRNA
reaction
product (1)
enzyme
treatment
25
°C
30 min
reaction
product (2)
ADPRC
3-azido-1-propanol
37
°C
30 min
reaction
product (3)
biotin-PEG4-DBCO
37
°C
60 min
a
PEG4-Biotin
AbTir-E/A
OH
O
O
O
O
P
P
O
+
RNA
OH OH
N
NH2
O -
O -
NN
N
N
NH2
AbTir
+
-
-
+
-
-+
-
-
(3)
+
-
-
+
-
-+
-
-
(3)
H2O
AbTir
AbTir-E/A
Arabidopsis mRNA
+
-
-
+
-
-+
-
-
(3)
E. coli mRNA
Streptavidin-HRP
EB stainning
Streptavidin-HRP
de
−8
−6
−4
−2
0
2
4
6
8
0 5 10 15 20
log2(CPM of regular RNA-Seq)
−log10(FDR)
0
10
20
30
−8
−6
−4
−2
0
2
4
6
8
0 5 10 15 20
log2(CPM of regular RNA-Seq)
−log10(FDR)
0
100
200
300
12 NAD-RNAs 1057 NAD-RNAs
AbTir-E/A
AbTir
f
AbTir
AbTir-E/A
-5.00
-4.00
-3.00
-2.00
-1.00
1.00
2.00
3.00
4.00
5.00
log2 Ratio of
NAD-RNA-Seq/regular RNA-Seq
sibD
sibE
sibC
yicG
leuA
yfgG
glvG
ilvL
yafD
ycbF
guaD
yfgC
amyA
Cho
ldtE
ykgS
yfcO
insG
yfjI
ybiY
Boronate gel
log2 Ratio of
NAD-RNA-Seq/regular RNA-Seq
log2 Ratio of
NAD-RNA-Seq/regular RNA-Seq
O
O
O
O
O
O
P
P
O
RNA
OH OH
O -
O -
NN
N
N
NH2
O
O
O
O
O
O
P
P
O
RNA
OH OH
O -
O -
NN
N
N
NH2
O
O
O
O
O
O
P
P
O
RNA
OH OH
O -
O -
NN
N
N
NH2
O
N
N
N
N
NH
2
O
OHO
OP
O
O
–
O
PO
N
+
O
OHOH
O
O
NH
2
O
–
RNA
N
N
N
N
NH
2
O
OHO
OP
O
O
–
O
PO
N
+
O
OHOH
O
O
NH
2
O
–
RNA
NN
+
N
-
N
N
N
N
NH
2
O
OHO
OP
O
O
–
O
PO O
OHOH
O
O
–
RNA
N
N
N
N
NH
2
O
OHO
OP
O
O
–
O
PO O
OHOH
O
O
–
RNA
Fig. 5 | AbTir deNAMs native NAD-RNAs. a Workflow to determine the deNAMing
activity of AbTir on cellular NAD-RNAs. In vitro transcribed NAD-RNA or total cel-
lular RNAs were pre-treated with AbTir or AbTir-E/A and then subjected to two
steps (ADPRC and SPAAC reactions) that convert NAD-RNAs to biotinylated RNAs.
In the presence of AbTir, NAD-RNAs are deNAMed and the subsequent biotinyla-
tion cannot occur. In the absence of AbTir, the ADPRC reaction replaces the
nicotinamideof the NAD cap with3-azido-1-propanol,which is then linked to biotin-
PEG4 in the SPAACreaction. bAn in vitrotranscribedNAD-RNA was used to test the
pipeline in (a). Thenumbers in brackets standfor the reaction products at different
stepsfrom the pipelinein (a). An APB gelwas used to resolvethe differentproducts,
which were visualized by ethidium bromide (EB) staining. To detect biotinylated
RNAs, a dot blot was performed using nylon N+membrane and biotin signals were
detected with streptavidin-HRP. cGel blots showing biotinylated RNAs after Ara-
bidopsis and E. coli RNAs went through the pipeline in (a). One microgram Arabi-
dopsis mRNA or 1 μgE. coli rRNA-depleted RNA was used for each assay. The RNA
products were resolved in 2% agarose gels and transferredto nylon N+membranes
for the detection of biotinsignals with streptavidin-HRP. d,eScatter plots showing
NAD-RNAs identified by SPAAC-NAD-Seq in E. coli RNA after AbTir (d)orAbTir-E/A
(e) pretreatment. The log2 ratio of CPM (Counts Per Million) between NAD-RNA-
Seq and regular RNA-Seq is plotted against the log2 CPMof regular RNA-Seq. The
carmine dots with different sizes represent enriched NAD-RNA-producing genes,
which was identified based on the criteria of “NAD-RNA-Seq/regular RNA-Seq ≥2
and FDR ≤0.05”.fHeatmap showing the levels of 20 known NAD-RNAs after AbTir
pretreatment. Fifteen (labeled with red fonts) of the 20 known NAD-RNAs were
identified as NAD-RNAs in our study, all of which were decapped by AbTir pre-
treatment. Four genes (sibD,amyA,ykgS,andyfjI) in grayfonts did not havecognate
genes in the E. coli BL21 (DE3) strain and another gene yfcO was not detected as an
NAD-RNA-producing gene. Source data are provided as a Source Data file.
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 7
Content courtesy of Springer Nature, terms of use apply. Rights reserved

AbTir
AbTir-E/A
grow at 37 °C
until OD600 ~0.4-0.8
IPTG induction
(0.1 mM, 0.5 mM, 1 mM, 5 mM)
incubate at 25°C
1. collect cells at different time points and
measure OD600
2. normalize OD600 to ~0.5, measure
endogenous NAD
+
levels
3. extract total RNA and measure NAD-RNA
content
a
time after IPTG induction (min)
AbTir
AbTir-E/A
time after IPTG induction (hour)
OD600
0
1
2
3
4
AbTir AbTir-E/A
0.1 mM 1 mM 0.5 mM 5 mM
IPTG concentrition
bcd
ef
g
h
** ** ** **
+-+-
i
−8
−6
−4
−2
0
2
4
6
8
0 5 10 15 20
−log10(FDR)
0
100
200
300
−8
−6
−4
−2
0
2
4
6
8
0 5 10 15 20
−8
−6
−4
−2
0
2
4
6
8
0 5 10 15 20
−8
−6
−4
−2
0
2
4
6
8
0 5 10 15 20
935 NAD-RNAs
914 NAD-RNAs
791 NAD-RNAs 1009 NAD-RNAs
AbTir-E/A IPTG-
AbTir-E/A IPTG+
AbTir IPTG+ AbTir IPTG-
@anti-MBP
CBB staining
AbTir
AbTir-E/A
24
114 138
66
39 68 44
27 79
14 546 38
19 98
24
AbTir
IPTG-
IPTG+
AbTir
-
E/A
IPTG+
IPTG-
P =0.0505
nuclease P1
++
--
AbTir AbTir-E/A
IPTG
++++
0
1
2
3
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−6
−4
−2
0
2
4
6
−6 −4 −2 0 2 4 6
●
●
●
all
NAD-RNAs
AbTir targets
absorbance (450 nm)
absorbance (450 nm)
log2(CPM of regular RNA-Seq)
log2 Ratio of
NAD-RNA-Seq/regular RNA-Seq
log2 Ratio of
NAD-RNA-seq/regular RNA-seq
log2Ratio of
NAD-RNA-Seq/regularR
NA-Seq
AbTir-E/A IPTG+
AbTir IPTG+
absorbance (450 nm)
024681012
0
1
2
3
4
*** ****
010 20 30 40 50 60 70 80 90
0
1
2
3
4
** ** **** ** ** **
N.S.
AbTir AbTir-E/A
N.S. N.S.
Fig. 6 | AbTir is a deNAMing enzyme in E. coli cells. a Diagram showing hetero-
logously expressing AbTiror AbTir-E/A in E. coli followed by evaluation ofits effects
on cell growth, endogenous NAD+,andNAD-RNAlevels.bMeasurement of OD600
values at the indicated time points after the addition with 1mM IPTG. Three bio-
logical replicates were measured at each time point. cLevels of NAD+in E. coli cells
expressing AbTir(orange) orAbTir-E/A(light blue)at 2 h after induction. Each bar
stands for a biological replicate. dLevels of NAD+in E. coli expressing AbTir
(orange) or AbTir-E/A (light blue) after induction with 0.1mM IPTG. Three biolo-
gical replicates were measured at each time point. eWestern blot to detect MBP-
tagged AbTir and AbTir-E/A proteins with anti-MBP, demonstrating that the
recombinant proteins werepresent at 15 min after IPTG induction.CBB, Coomassie
Brilliant Blue. fQuantification of NAD cap content in total RNA isolated from E. coli
cells expressingAbTir or AbTir-E/Aafter 15 min IPTG induction.Nuclease P1 (+) was
used to digest the total RNA to release the NAD cap. Samples treated with DEPC
water (-) were used as negative controls. gScatter plots showing NAD-RNAs iden-
tified by SPAAC-NAD-Seq in E. coli expressing AbTir or AbTir-E/A with or without
IPTG induction. The log2 ratioof CPM between NAD-RNA-Seqand regular RNA-Seq
is plotted against the log
2
(CPM) of regular RNA-Seq. The labeled dots (carmine
color) represent the identified NAD-RNAs. hVenn diagram showing the overlap of
NAD-RNAs identified in E. coli expressing AbTir or AbTir-E/A with or without IPTG
induction. Bold numbers represent NAD-RNAs identified in E. coli expressing AbTir
that were also identified in the other three datasets. Red numbers represent the
NAD-RNA-producing genes that were specifically identified in E. coli without AbTir
expression. iComparison of NAD-RNA levels between E. coli expressing AbTir and
AbTir-E/A. NAD-RNA levels are defined as the log2 ratio of NAD-RNA-Seq/regular
RNA-Seq. Red points represent NAD-RNAs identified in E. coli expressing AbTir-E/A.
Blue points represent the NAD-RNAs thathave significantly lowerenrichment levels
in E. coli cells expressing AbTir than in E. coli expressing AbTir-E/A. For (b,d,f),
average values from three biological replicates are shown with error bars denoting
mean ± SD. [**] P≤0.01; [*] P≤0.05; N.S., not significant (calculated by the non-
parametric Mann-Whitney U-test, two-sided). Exact P-values and source data are
provided as a Source Data file.
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 8
Content courtesy of Springer Nature, terms of use apply. Rights reserved

IPTG), wecollected AbTir-and AbTir-E/A-expressing cells and isolated
total RNAs. After removing free NAD+, we performed nuclease P1
digestion on the total RNA and quantified the NAD cap content using
the NAD-capQ assay. We observed a significant, albeit slight, reduction
inNADcapcontentintotalRNAfromE. coli cells expressing AbTir
compared to AbTir-E/A (Fig. 6f). This suggests a global reduction in
NAD-RNA levels in vivo after AbTir expression.
To determine changes in specific NAD-RNA transcripts, SPAAC-
NAD-Seq7,15 was carried out with the same total RNAs from E. coli cells
expressing AbTir and AbTir-E/A (n = 3 for each group).AfterrRNA
depletion, the retained RNA was subjected to treatments that con-
verted NAD-RNAs to biotin-RNAs for sequencing (referred to as “NAD-
RNA-Seq”). A portion of the rRNA-depleted RNA was used for direct
RNA sequencing (referred to as “regular RNA-Seq”) as a control. The
sequencing libraries were of high reproducibility, according to clus-
tering analysis (Supplementary Fig. 14). We identified a total of 791
NAD-RNA-producing genes in E. coli cells expressing AbTir after IPTG
induction, which is fewer than the number identified in E. coli cells
without IPTG induction or in E. coli cells expressing AbTir-E/A (Fig. 6g;
Supplementary Data 3–6). Overlap analysis revealed that 767 (~97%) of
the NAD-RNA-producing genes identified in E. coli cells expressing
AbTir were consistently identified in the other three datasets,
demonstrating the accuracy and effectiveness of SPAAC-NAD-Seq
(Fig. 6h). A total of 547 NAD-RNA-producing genes were specifically
identified in E. coli cells without AbTir expression (Fig. 6h), suggesting
that they are the possible decapping targets of AbTir in vivo. Using
more stringent criterion of at least a 2-fold change and P≤0.05, we
identified 78 NAD-RNA-producing genes that had higher levels of NAD-
RNAs in E. coli cells expressing AbTir-E/A than in cells expressing AbTir
(Fig. 6i;Supplementary Fig.15; Supplementary Data 5). These genes are
considered deNAMing targets of AbTir in vivo. Functional enrichment
analysis showed that these AbTir-sensitive NAD-RNA-producing genes
are most strongly enriched in GO terms including “molecule transport
process”and “oxidoreductase activity”.Thesefindings indicate that
AbTir, as a deNAMing enzyme, preferentially targets a subset of NAD-
RNAs in E. coli cells. However, it should be noted that the number of
AbTir deNAMing targets is likely underestimated because we collected
cells for RNA isolation only at 15min after IPTG induction in order to
avoid the disturbance of free NAD+.
TcpA, a TIR domain-containing protein from archaea, also
exhibits deNAMing activity
Recent studies have demonstrated the presence of NAD-RNAs in
archaeal model organisms, with the highest NAD cap concentration
reported so far9,16, thereby extending the existence of NAD-RNAs to all
three domains of life. Given the widespread occurrence of NAD-RNAs
and the conservation of TIR domains across various phyla59,60,we
hypothesize that the deNAMing capacity of TIR domain might be
evolutionarily conserved. To explore this possibility, we investigated
the deNAMing ability of several TIR homologs from archaea to plants
and humans (Fig. 7a). We first expressed two TIR domain-containing
proteins, TcpA and TcpO from archaeal species Theionarchaea
archaeon and Methanobrevibacter olleyae,respectively,inE. coli.
Strikingly, TcpA depleted free NAD+as expected35 and cleaved the RNA
NAD cap, similar to AbTir (Fig. 7b, c). Both activities were abolished in
the putative catalytic mutant (TcpA-E/A) (Fig. 7b, c). However, we did
not observe any decapping activity for TcpO, despite its apparent
NADase hydrolytic activity35 (Fig. 7b, c), suggesting that not all TIR
domain-containing proteins possess the NAD-RNA deNAMing activity.
Additionally, TIR domains are present in a wide range of microbial
genomes34,36, and certain microbial TIR-domain NADaseshave been co-
opted as virulence factors34,37,61. For instance, HopAM1, a TIR domain-
containing effector protein encoded by the plant pathogen Pseudo-
monas syringae DC3000, has been shown recently to suppress plant
immunity by hydrolyzing NAD+and producing a novel cADPR variant
(v2-cADPR or 3′-cADPR) in plant cells34,37. With the purified protein
(Fig. 7d), we confirmed the NADase activity of HopAM1, which was
dependent on the putative catalytic Glu/E residue (E191) (Fig. 7e).
However, we did not observe the decapping activity of HopAM1 on
NAD-RNAs,evenwithhighconcentrationsoftheHopAM1protein
applied (Fig. 7f). Given the cyclization site of 3′-cADPR, it is theoreti-
callyimpossibletogeneratea3′-cADPR-RNA, because the 3′-OH on the
ribose of the cap adenosine is occupied by the RNA body. It is still
unknown if there are other TIR domain-containing effectors having
decapping ability on NAD-RNAs, which might enzymatically subvert
plant immune systems.
The classical human TIR NADase SARM1, which can hydrolyze
free NAD+efficiently39,51 (Supplementary Fig. 16a, b), was also exam-
ined and proven incapable of cleaving the RNA NAD cap (Supple-
mentary Fig. 16c). To further extend our testing to plants, we selected
several TIR domain proteins, including BdTIR from Brachypodium
distachyon, OsTIR from Oryza sativa, the TIR domain of L7 from
Linum usitatissimum, and the TIR domains of RPS4 (resistance to
Pseudomonas syringae 4) and RBA1 (Response to HopBA1) from Ara-
bidopsis thaliana, for expression in E. coli.However,consistentwith
previous reports, the plant TIR domains did not deplete free NAD+in
our in vitro NADase assay due to their weak NADase activities.
Moreover, some of them showed strong nuclease activities (i.e., 2′,3′-
cAMP and 2′,3′-cGMP synthetases) as reported40 (Supplementary
Fig. 16d), which hampered our assays to examine the deNAMing
activity with in vitro transcribed NAD-RNAs. Although more investi-
gation is definitely required, our results showed that the deNAMing
activity of TIR domain-containing proteins might be evolutionarily
conserved, at least in bacteria and archaea.
Discussion
The existence of NAD-RNAs has been well-established in various
organisms5–16, uncovering a potential connection between NAD+
metabolism and gene expression. Identification of the enzymatic
machinery for adding or removing the NAD cap will help understand
the biological functions of NAD-RNAs. Increasing studies over the past
years have defined TIR domain-containing proteins as NADase
enzymes that hydrolyze free NAD+in both prokaryotic and eukaryotic
models. Here, our initial screening of putative NAD cap decapping
enzymes led to the identification of TIR domain-containing proteins as
NAD-RNA deNAMing enzymes, furthering our understanding of TIR
domain’s functionality. We demonstrated that AbTir, one of the bac-
terial TIR domain-containing proteins, can cleave in vitro synthesized
and native NAD-RNAs using a combination of methods (Figs. 1and 2),
and v-cADPR-RNA was identified as the potential RNA product after
cleavage (Fig. 3). However, whether v-cADPR-RNA exists in vivo, and if
so, whether it is functional, is so far unknown. Our current study
showed that NAD-RNAs canbe first converted into cyclic ADPR-RNA by
TIR domain-containing proteins, which can be further converted into
p-RNA by Nudix or DXO/Rai1 decapping enzymes (Fig. 4c). This sug-
gests the potential existence of a two-step NAD-RNA decapping
strategy in vivo, which would be specifictoNAD-RNAsduetothe
specificity of the deNAMing enzymes towards NAD-RNAs.
With our current results, only some of the bacterial and archaeal
TIR domain-containing proteins showed deNAMing activity on NAD-
RNA. It is unknown what features allow these TIR domain-containing
proteins to possess such an activity. TIR domain-containing proteins
are composed of versatile modules with distinct domain architectures
across plants, animals, and prokaryotes60. However, we did not find
any common architecture apart from the TIR domain among AbTir,
BtpA, PdTir, and TcpA. We attempted to examine structural models of
the investigated TIR domain proteins from both bacteria and Archaea.
Interestingly, we found that all TIR domain-containing proteins with
decapping activity (AbTir, BtpA, PdTir, and TcpA) tend to possess
positive charges on the protein surface near the catalytic site, whereas
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 9
Content courtesy of Springer Nature, terms of use apply. Rights reserved

TcpO -- the archaeal TIR domain protein without deNAMing activity --
exhibits a large block of negative charges (Supplementary Fig. 17).
Considering the negative charges carried by RNA molecules, it is
plausible that the TIR domain-containing proteins with deNAMing
activity might have an affinity for RNA. However, the robust negative
charges present on the TcpO surface may act to repel RNA molecules.
It should be noted that biochemical experiments with purified proteins
cannot recapitulate the true biologically active conformation of some
TIR domain-containing proteins, especially for those from plants that
have weak NADase activity and usually require an oligomerized com-
plex to function. Thus, additional studies willbe necessary to address
whether TIR domain proteins in plants and animals have NAD-RNA
deNAMing activity.
A plethora of recent studies underscored the importance of the
NADase activity of TIR domain-containing proteins in immunity and
proposed that NAD+depletion or the generation of cleavage products
might be a signaling event triggering downstream immune responses
that lead to cell death38,40,41,47,56. Interestingly, nuclear localization of
some plant TIR domain-containing proteins is required for disease
resistance62, which implies that TIR domain proteins might mediate
gene expression regulation in the nucleus. Our study uncovered a
novel enzymatic TIR domain function that can potentially reprogram
the NAD-RNA transcriptome and raised the possibility that some TIR
domain-containing proteins could regulate gene expression through
NAD-RNA deNAMing.
Methods
Bacterial strains and culture media
E. coli DH5αand BL21 (DE3) strains used for plasmid construction and
protein expression, respectively, were grown in Luria-Bertani (LB)
medium amended with selective antibiotics at 37°C. Ampicillin was
included at a concentration of 50 µg/mL.
Plasmids construction and PCR-mediated site-directed
mutagenesis
Double-tagged (N-terminal MBP and C-terminal 6×His tags) TIR
domain-containing proteins (full length or TIR domain-only) from
bacteria, archaea, and plants were cloned into a modified pMAL-C2X
empty backbone to prepare prokaryotic expression plasmids.
Recombinant plasmids include MBP-AbTir-6×His (1-268aa), MBP-BtTir-
6×His (1-287aa), MBP-TcpCTIR−6×His (173-307aa), MBP-BtpA-6×His (1-
275aa), MBP-TcpF-6×His (1-274aa), MBP-PdTir-6×His (1-299aa) and
MBP-TirS-6×His (1-280aa) from Bacteria, MBP-TcpA-6×His (1-327aa)
and MBP-TcpO-6×His (1-341aa) from archaea,RPS4TIR+80aa (1-280aa) and
RBA1 (1-362aa) from Arabidopsis, and BdTir (1-224aa) and OsTIR (1-
196aa) from Brachypodium distachyon and Oryza sativa,respectively.
DNA sequences optimized for prokaryotic protein expression were
synthesized at Twist Bioscience and used as DNA templates for PCR
amplification. The PCR products were gel-purified and confirmed by
Sanger sequencing, and then ligated into the modified pMAL-C2X
vector by Gibson Assembly cloning. PCR-mediated site-directed
NAD-RNA
cleavage prduct
ppp-RNA
absorbance (450 nm)
elution buffer
****
Tcp A
Tcp A - E / A
**
Tcp O
0
0.5
1.0
1.5
2.0
2.5
3.0
TcpO-E/A
**
HopAM1
HopAM1-E/A
250-kDa
70-kDa
55-kDa
35-kDa
100-kDa
15-kDa
0
0.5
1.0
1.5
2.0
2.5
3.0
elution buffer
AbTir
HopAM1
HopAM1-E/A
absorbance (450 nm)
**
N.S.
**
ab
c
de f
Bacteria Annimal
Plant
Archaea Fungus
Tcp A
Tcp O
AbTir
BtTir
BtpA
Tcp C
BdTIR
OsTIR
RPS4
RBA1
SNC1
HopAM1 hSARM1
Tcp F
TirS
PdTir
NAD-RNA
cleavage
prduct
hopAM1
TcpO-E/A
TcpO
TcpA - E / A
ADPR-RNA
NAD-RNA
-+
++++++
-
-
protein
TcpA
-----
--
20 μM
10 μM
5 μM
NAD-RNA
+++++
-
protein
2 μM
+++
AbTir
hopAM1-E/A
40
μM
HopAM1-E/A
HopAM1
NAD-RNA
++++
-
protein
AbTir
Fig. 7 | Evaluation of the NADase and deNAMing acitivties of TIR domain-
containing proteins in other organisms. a The presence of typical TIR domain-
containing proteins in all three domains of life. Red font labeled TIR domain-
containing proteins have been confirmed to exhibit deNAMing activities in our
study. bNADase activities of TcpA and TcpO from archaea. cAn APB gel showing
the RNA species after an in vitro transcribed NAD-RNA was incubated with TcpA or
TcpO. In vitrotranscribed ADPR-RNA and NAD-RNA were included as makers. dAn
SDS-PAGE gel showing the recombinant HopAM1 and HopAM1-E/A proteins pur-
ified from BL21 E. coli cells. The proteins were resolved in a 15% SDS-PAGE gel and
visualized by Coomassie BrilliantBlue staining. eNADase activities of HopAM1. The
catalytic mutant HopAM1-E/A was included as a negative control. AbTir was inclu-
ded as a positive control. fAPB gels showing the RNA species after reactions in
which different concentrations of HopAM1 were incubated with an in vitro tran-
scribed NAD-RNA. The reaction with AbTir was included as a positive control. The
y-axes in (b,e) represent relative NAD+levels, which were measured with the NAD/
NADH Quantitation Kit by monitoring the absorption values at 450 nm. Error bars
represent mean ± SD, which was calculated from five independent experiments
(n=5); [**]P≤0.01; N.S., not significant (calculated by the non-parametric Mann-
WhitneyU-test, two-sided). ExactP-values and sourcedata are providedas a Source
Data file.
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 10
Content courtesy of Springer Nature, terms of use apply. Rights reserved

mutagenesis was performed to produce the catalytic mutants,
including MBP-AbTir-E/A-6×His, MBP-BtpA-E/A-6×His, MBP-PdTir-E/A-
6×His, MBP-TcpA-E/A-6×His, and MBP-TcpO-E/A-6×His. All primers
involved in constructing the above plasmids and generating the point
mutations are listed in Supplementary Table 1.
Recombinant protein expression and purification
The plasmids containing the TIR domains with the N-terminal MBP tag
and the C-terminal 6 × His tag were transformed into competent E. coli
BL21 cells. Single colonies were cultured in 1 mL LB containing 50μg/
mL ampicillin and grown overnight at 37 °C. These cultures were used
toinoculate400mLofthesamemediumat1:100ratio.Thelarge
cultures were grown at 37 °C for 3-4 h until an OD value at 600 nm
(OD600) of 0.4–0.6. Then, 0.2 mM isopropyl β-D-1-thiogalactopyr-
anoside (IPTG; Thermo Fisher Scientific, 15529019) was added to
induceproteinexpression,andthecellsweregrownat16°Cfor∼16 h.
The bacterial cells were collected by centrifugation at 5000 g for 15 min
at 4 °C. For protein purification, cell pellets were first resuspended in
20 mL of lysis buffer (100 mM Tris-HCl, pH 7.5; 500 mM NaCl; 5 mM 2-
mercaptoethanol; 5% [v/v] glycerol; 20 mM imidazole; 0.1% [v/v]
CA630) supplied with 1 mM protease inhibitor PMSF and 1× EDTA-free
protease inhibitor cocktail. The cells were lysed by sonication, and the
insoluble cell debris was removed by centrifugation at 10,000 g for
40 min. The recombinant proteins were purified with Profinity™IMAC
Resin, Ni-charged (Bio-Rad, 1560133). The proteins were eluted with the
elution buffer (50 mM Tris-HCl, pH 7.5; 500 mM NaCl; 5 mM 2-mer-
captoethanol; 5% [v/v] glycerol; 400mMimidazole;0.1% [v/v]CA630)
and the concentrations were determined using the Bradford assay (Bio-
Rad, Quick Start™Bradford 1×Dye Reagent #5000205).
Measurements of NAD+levels in E. coli cells
The recombinant constructs of TIR domain proteins, as well as their
corresponding catalytic E/A mutants, were transformed into compe-
tent E. coli BL21 cells. After confirmation by Sanger sequencing, single
colonies were then cultured at 37 °C until they reached an OD600 of
approximately 0.4–0.8, when IPTG was added to induce protein
expression. The cultures at 2 h after IPTG induction were adjusted to
OD600 = 0.5 ± 0.05, and the pellets from 5 00 μL of culture suspension
were lysed with 200 μL 0.5 M perchloric acid (HClO
4
). Samples were
immediately placed on ice for 10min and centrifuged at 12000g for
10 min. 180 μL supernatant from each sample was transferred to a new
tube, and 67 μLof3MK
2
CO
3
was added. The reactions were placed on
ice for another 10 min and centrifuged at 12000 g for 10min. After
centrifugation, 50 μL supernatant containing the extracted metabo-
lites from each sample was mixed well with the Master Reaction Mix,
which is a mixture of 2 μL NAD Cycling Enzyme Mix and 98μLNAD
Cycling Buffer from the NAD/NADH Quantitation Kit (Sigma-Aldrich,
MAK037). The mixture was incubated for 5 min under dark condition
at room temperature, allowing the reduction of NAD+to NADH. Finally,
10 μL NADH developer from the NAD/NADH Quantitation Kit was
added into each sample to react with NADH and a product that absorbs
light at a wavelength of 450nm was generated. The samples were
transferred into a 96-well plate and measured for absorbance at
450 nm by spectrophotometry.
RNA synthesis by in vitro transcription
A single-stranded DNA sequenceT7ɸ2.5-A-32 (5′-CAGTAATACGACTCA
CTATTAGGCCTCTCGCTCTGCTGGGTGTGCGCTTGC-3′), which con-
tains a T7 ɸ2.5 promoter at the 5′-end and a unique adenosine at the
transcription start site, and its reverse complementary sequence (5′-
GCAAGCGCACACCCAGCAGAGCGAGAGGCCTAATAGTGAGTCGTATT
ACTG-3′) were synthesized and annealed with each other to get the
double-stranded DNA (dsDNA) template. In vitro transcription was
carried out at 37 °C overnight with the reaction (100 μL) containing
2μg of the dsDNA template, 1 × T7 polymerase buffer (New England
Biolabs), 1 mM CTP, 1 mM GTP, 1 mM UTP, and 1 mM ATP (for ppp-
RNA) or NAD+(for NAD-capped RNA) or FAD (for FAD-capped RNA) or
dpCoA (for dpCoA-capped RNA) or ADPR (for ADPR-capped RNA) or
Ap
4
A(forAp
4
A-capped RNA), and 1 U/μL T7 RNA polymerase (New
England Biolabs; M0251S). The RNA products were treated with DNase
I (Roche) at 0.2 U/μL at 37 °C for 30 min, extracted by phenol/
chloroform (5:1, pH 4.5), and then precipitated with ethanol. Unin-
corporated nucleotides were removed using Micro Bio-Spin P-30 Gel
Columns (Bio-Rad) or by performing gel recovery after acryloylami-
nophenyl boronic acid affinity gel electrophoresis.
Acryloylaminophenyl boronic acid (APB) affinity gel
electrophoresis
APB affinity electrophoresis was employed for the purification and
analysis of 5′-cap-modified RNAs42.TopreparetheAPBaffinity gel,
80 mg 3-acrylamidophenylboronic acid (Boron Molecular, CAS:
99349-68-5) was dissolved into a mixture of the polyacrylamide solu-
tion (30 mL; including 1.5 mL 5 × TBE, 6.3 g urea, 5.76 mL 40% Acryla-
mide/Bis Solution) by rotating for 10 min. The solution was
polymerized with APS (0.1% final concentration) and TEMED (0.1% final
concentration). 0.5 × TBE was used as the gel running buffer for the
APB gel. For the preparation of pure 5′-cap-modified RNAs, the in vitro
transcribed RNA product was added to an equal volume of 2× RNA
loading dye (80% Formamide, 0.1% Xylene FF, 0.1% Brophenol Blue)
and denatured at 65 °C for 5 min. After denaturing, the sample was
immediately cooled down on ice for gel loading. The modified RNAs
were collected by excising the gel and then eluted from the gel slice.
For the analysis of the RNA products of various decapping reactions,
the reactions were stopped by adding an equal volume of 2× RNA
loading dye and RNAs were denatured at 65 °C for 5 min. After APB gel
electrophoresis, the gel was stained with ethidium bromide for 2min
on a shaker and visualized in the Molecular Imager® Gel Doc™XR
System.
NADase and deNAMing/decapping assays
For NADase assays, the 20 μLreactioncontained20μMNAD
+,10μM
purified proteins in NADase reaction buffer (50 mM Tris-HCl, pH 8.0;
100 mM NaCl; 20 mM MgCl
2
), and 20% PEG6000. After incubation at
25 °C for various durations as specified in Results, 30 μL NAD/NADH
extraction buffer from the NAD/NADH Quantitation Kit (Sigma-
Aldrich, MAK037) was added to the reaction for measuring the NAD+
content. For RNA decapping assays, 500ng in vitro transcribed RNAs
(32-nt) capped with NAD, FAD, dpCoA, ADPR, or Ap
4
Awereincubated
with the purified proteins (10 μM) in decapping buffer (50 mM Tris-
HCl, pH 8.0; 100 mM NaCl; 20 mM MgCl
2
; 20% PEG6000) at 25 °C for
various durations as specified in Results. The reactions were mixed
with an equal volume of 2 × RNA loading dye and RNAs weredenatured
at 65 °C for 5 min. The RNAs were resolved by running a denaturing 15%
(w/v) APB gel with 0.5 × TBE as the running buffer and visualized by
ethidium bromide staining.
NAD cap detection and measurement with the NAD-
capQ method
To measure the amount of the NAD cap before and after AbTir treat-
ment, the NAD-capQ method was performed aspreviously described43.
Briefly, in vitro transcribed NAD-RNA (1 μg) or total RNA (500 μg)
isolated from E. coli (stationary phase) or Arabidopsis (12-day-old
seedlings) weresubjected to AbTir or AbTir-E/A treatment at 25°C for
16 h. Then, the RNA products were extracted with phenol/chloroform
(pH 4.5),precipitated, and subjected to nuclease P1 digestionin a 20 μL
reaction with 1 × nuclease P1 Reaction Buffer and 10 U/μLnucleaseP1
(New England Biolabs; M0660S) at 37°C for 2h. Before the AbTir
cleavage reaction, the isolated total RNA was treated with NAP-10
columns to get rid of any residual free NAD+. Following digestion with
nuclease P1, 30 μL of NAD/NADH Extraction Buffer from the NAD/
Article https://doi.org/10.1038/s41467-024-46499-y
Nature Communications | (2024) 15:2261 11
Content courtesy of Springer Nature, terms of use apply. Rights reserved

NADH Quantification Kit (Sigma-Aldrich, MAK037) was added to each
sample. The 5 0 μL reactions were then used for the subsequent
enzymatic cycling reaction and the colorimetric assay by following the
manufacturer’s protocol from the NAD/NADH Quantitation Kit. The
same RNA samples treated with the AbTir-E/A catalytic mutant served
asthenegativecontrols.
HPLC-MSanalysisofvariousmetabolites
Liquid chromatography was performed on a Nexera XR 40 series HPLC
(Shimadzu) using a Synergi 4 µM Fusion-RP 80 Å 150 × 2 mm column
(Phenomenex). The column temperature was kept at 40 °C, and the
sample tray was maintained at 4 °C. 10 μL samples were injected at a
flow rate of 0.2ml/min with 10 mM ammonium formate (pH 4.2) and
methanol as mobile phases A and B, respectively. Metabolites were
eluted using the following gradient: 0–8min,8–90% B; 8–10 min, 90%
B; 10–10.1 min, 90–8% B; 10.1–20 min, 8% B. The LCMS-8060 triple
quadrupole mass spectrometer with electro spray ionization (Shi-
madzu) was operated in the positive mode. Scheduled multiple reac-
tion monitoring (MRM) was employed to monitor analyte parent ions
to product ion formation. MRM conditions were optimized using
authentic standard chemicals including NAM ([M + H]+123.00 > 80.00,
123.00 > 78.00, 123.00 > 53.00), NAD+([M + H]+664.00 > 136.00,
664.00 > 427.90, 664.00 > 523.95), cADPR ([M + H]+541.80 > 136.15,
541.80 > 427.90, 541.80 > 347.90), and ADPR ([M + H]+559.80 > 136.05,
559.80 > 427.95, 559.80 > 347.95). Data acquisition was performed
using the LabSolutions LCMS v5.97 software, and data processing was
carried out using LabSolutions Postrun (both Shimadzu). Metabolite
products were quantified by scheduled MRM peak integration using
calibration curves of standard chemicals.
Cellular RNA isolation from E. coli BL21 strains and ribosomal
RNA (rRNA) depletion
The culture of E. coli BL21 after overnight growth was diluted into fresh
medium ata 1:100 ratio and the new culture was grown until an OD600
of 3.0 (the stationary phase). Isolation of total RNA was performed
according to a published method63. In brief, cell pellets were collected
by centrifugation at 6000 g for 10 min at 4 °C, and immediately put on
ice for re-suspension by adding 20 mL TES buffer (10 mM Tris-HCl,
pH7.5; 10 mM EDTA; 0.5% (w/v) SDS). The re-suspended cells were
mixed with an equal volume of phenol/chloroform (5:1, pH 4.5) and
incubated for 5–10 min at 65 °C with vigorous shaking. The cells were
vortexed for 5min at room temperature and centrifuged for 10 min at
12,000g at 4 °C. The upper phase wascollected and mixed with20 mL
of phenol/chloroform (5:1, pH 4.5) again. After another round of cen-
trifugation (12,000g, 4 °C), the upper phase was mixed with 10 ml
chloroform. Then, the upper phase was gently collected after 10 min of
centrifugation (12,000 g at 4 °C) and mixed with isopropanol at a 1:1
ratio. The mixture was incubated for 15min at room temperature for
RNA precipitation. Total RNA was pelleted by centrifugation (12,000 g
at 4 °C) for 30min, washed twice with 75% ethanol, and dissolved in
DEPC H
2
O.
Depletion of the 16S and 23S rRNAs was performed using the
MICROBExpress™Bacterial mRNA Enrichment Kit (Invitrogen,
AM1905) according to manufacturer’s instructions. The resulting
rRNA-depleted RNA contains mRNA, tRNA, 5S rRNA, and other small
RNAs and was used for subsequent experiments.
Identification of NAD-capped RNAs by SPAAC-NAD-Seq
The rRNA-depleted RNA from 10 μgE. coli total RNA was used for
preparing the SPAAC-NAD-Seq libraries according to ref. 15.Atthe
same time, 100ng of rRNA-depleted RNAs were used for regular RNA-
Seq library construction as a control for NAD-RNA identification.
NEBNext® Ultra™II RNA Library Prep Kit for Illumina® (New England
Biolabs, E7770) was employed to generate libraries for regular RNA-
Seq and SPAAC-NAD-Seq with three biological replicates. After
sequencing, the clean data were first examined for the library quality
with the FastQC program (https://github.com/s-andrews/FastQC). The
data after quality check were subjected to a series of analyses,
including low-quality nucleotide trimming, reads alignment, frag-
ments counting, and differential gene expression analysis. In brief, the
3′-end low-quality nucleotides were trimmed with trim_galore (https://
www.bioinformatics.babraham.ac.uk/). The remaining reads longer
than 100-bp were mapped to the E. coli BL21 genome (https://www.
genome.jp/kegg-bin/show_organism?org=ebl) using bowtie2 with
default parameters. Next, HTSeq-count64 was used to count reads that
mapped to the genic regions. NAD-RNA-producing genes were iden-
tified with ratio of counts per million(CPM) between NAD-RNA-Seq and
regular RNA-Seq ≥2andFDR≤0.05 using edgeR65.
Structural modeling
AlphaFold266 was used to predict the TIR domain structures of TcpA,
BtpA, and TcpO. The predicted monomers of TcpA, BtpA, and TcpO
were individually aligned onto the TIR domains of AbTir NADase
asymmetric dimmer (7UXU). The PdTIR (3H16) protomer was also
aligned onto the TIR domain of the AbTir NADase asymmetricdimmer.
Since both the PdTIR structure and the predicted TIR structures of
TcpA, BtpA and TcpO are in their inactive states, the BB-loops inthese
proteins present a closed conformation (Supplementary Fig. 17).
Electrostatic potentials on the surface of proteins were visualized by
PyMOL (v2.5) and color-coded, with red indicating negative charges,
blue denoting positive charges, and white representing neutral
charges.
Statistics and reproducibility
Statistical analyses to examine significant differences and result
visualization were performed with the GraphPad Prism 9 software.
Significance was determined by p< 0.05 [*] or p< 0.01 [**]. All gel
blots/plots were performed at least two times except for those
in Fig. 5b.
Reporting summary
Further information on research design is available in the Nature
Portfolio Reporting Summary linked to this article.
Data availability
The clean data of regular RNA-Seq and NAD-RNA-Seqin this study have
been deposited in National Center for Biotechnology Information
(NCBI) (BioProject database, accession code PRJNA1061094). All plas-
mids and strains supporting the finding of this study are availabl e from
the corresponding author uponrequest. Source data areprovided with
this paper.
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Acknowledgements
We would like to thank Dr. Liang Tong from Columbia University for his
helpful discussions and suggestions. We also thank Dr. Ming Guo from
University of Nebraska-Lincoln for generously sharing the expression
plasmids of hopAM1 and hopAM1-E/A. This work was supported by
grants from National Institutes of Health (NIH, GM061146) and Ministry of
Science and Technology of the People’s Republic of China (Chinese
Ministry of Science and Technology, 2023YFC3402200) to X.C.
Author contributions
X.W. and X.C. designed the project; X.W., D.Y., J.Y., H.H., R.H., Z. A., Q.C.
and J.C. conducted laboratory experiments; X.W. performed the RNA-
Seq data analysis and interpretation; X.C. supervised the project; X.W.
and X.C. wrote the manuscript. All authors reviewed and revised the
manuscript.
Competing interests
The authors declare no competing interests.
Additional information
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