Natural occurrence of 2′,5′-linked heteronucleotides in marine sponges

Article abstract of DOI:10.3390/md8020235

2′,5′-oligoadenylate synthetases (OAS) as a component of mammalian interferon-induced antiviral enzymatic system catalyze the oligomerization of cellular ATP into 2′,5′-linked oligoadenylates (2-5A). Though vertebrate OASs have been characterized as 2′-nucleotidyl transferases under in vitro conditions, the natural occurrence of 2′,5′-oligonucleotides other than 2-5A has never been demonstrated. Here we have demonstrated that OASs from the marine sponges Thenea muricata and Chondrilla nucula are able to catalyze in vivo synthesis of 2-5A as well as the synthesis of a series 2′,5′-linked heteronucleotides which accompanied high levels of 2′,5′-diadenylates. In dephosphorylated perchloric acid extracts of the sponges, these heteronucleotides were identified as A2′p5′G, A2′p5′U, A2′p5′C, G2′p5′A and G2′p5′U. The natural occurrence of 2′-adenylated NAD⁺ was also detected. In vitro assays demonstrated that besides ATP, GTP was a good substrate for the sponge OAS, especially for OAS from C. nucula. Pyrimidine nucleotides UTP and CTP were also used as substrates for oligomerization, giving 2′,5′-linked homo-oligomers. These data refer to the substrate specificity of sponge OASs that is remarkably different from that of vertebrate OASs. Further studies of OASs from sponges may help to elucidate evolutionary and functional aspects of OASs as proteins of the nucleotidyltransferase family.

Full text of DOI:10.3390/md8020235

Mar. Drugs 2010, 8, 235-254; doi:10.3390/md8020235
OPEN ACCESS
Marine Drugs
ISSN 1660-3397
www.mdpi.com/journal/marinedrugs
Article
Natural Occurrence of 2′,5′-Linked Heteronucleotides in Marine
Sponges
Annika Lopp, Tõnu Reintamm, Anne Kuusksalu, Indrek Tammiste, Arno Pihlak and
Merike Kelve *
Department of Gene Technology, Tallinn University of Technology, Akadeemia tee 15, 12618 Tallinn,
Estonia; E-Mails: annika.lopp@ttu.ee (A.L.); tonu.reintamm@gmail.com (T.R.);
anne.kuusksalu@ttu.ee (A.K.); indrek.tammiste@ttu.ee (I.T.); arpihlak@gmail.com (A.P.)
* Author to whom correspondence should be addressed; E-Mail: merike.kelve@ttu.ee;
Tel.: +372-6204432; Fax: +372-6204401.
Received: 23 December 2009; in revised form: 18 January 2010 / Accepted: 1 February 2010 /
Published: 2 February 2010
Abstract: 2′,5′-oligoadenylate synthetases (OAS) as a component of mammalian
interferon-induced antiviral enzymatic system catalyze the oligomerization of cellular ATP
into 2′,5′-linked oligoadenylates (2-5A). Though vertebrate OASs have been characterized
as 2′-nucleotidyl transferases under in vitro conditions, the natural occurrence of
2′,5′-oligonucleotides other than 2-5A has never been demonstrated. Here we have
demonstrated that OASs from the marine sponges Thenea muricata and Chondrilla nucula
are able to catalyze in vivo synthesis of 2-5A as well as the synthesis of a series 2′,5′-linked
heteronucleotides which accompanied high levels of 2′,5′-diadenylates. In
dephosphorylated perchloric acid extracts of the sponges, these heteronucleotides were
identified as A2′p5′G, A2′p5′U, A2′p5′C, G2′p5′A and G2′p5′U. The natural occurrence of
2′-adenylated NAD+ was also detected. In vitro assays demonstrated that besides ATP,
GTP was a good substrate for the sponge OAS, especially for OAS from C. nucula.
Pyrimidine nucleotides UTP and CTP were also used as substrates for oligomerization,
giving 2′,5′-linked homo-oligomers. These data refer to the substrate specificity of sponge
OASs that is remarkably different from that of vertebrate OASs. Further studies of OASs
from sponges may help to elucidate evolutionary and functional aspects of OASs as
proteins of the nucleotidyltransferase family.
Keywords: 2′,5′-oligoadenylate synthetase; OAS; marine sponge; 2′,5′-linked
heteronucleotides; natural products

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1. Introduction
2′,5′-oligoadenylate synthetases (OAS; EC 2.7.7.-) were discovered as a part of the interferon
antiviral pathway in mammals [1]. When activated by double-stranded RNA, the enzymes catalyze the
oligomerization of cellular ATP into 2′,5′-linked oligoadenylates (2-5A) with the general formula
pppA(2′p5′A)n, where n ≥ 1. Currently, the only known function of 2-5A is to bind and activate a
latent endoribonuclease (RNase L) that degrades single-stranded RNA of viral and cellular origin
[2–4]. These enzymes together with 2′-phosphodiesterase which degrades 2-5A, constitute an RNA
degradation pathway referred to as the 2-5A system [5]. In addition to functioning in the antiviral host
defense, OAS as a component of 2-5A system has also been implicated in the regulation of other
cellular processes, such as cell growth, differentiation and apoptosis [6–9]. Besides, biological roles
for OAS proteins distinct from activation of the RNase L pathway have been suggested. Ghosh et al.
have shown that a specific isozyme of OAS can induce apoptosis which is not mediated by the
endoribonuclease [10]. OAS may regulate gene transcription and thereby have additional effects on
cytokine receptor signal transduction pathways [11]. Physiological concentrations of 2-5A can be
involved in the regulation of gene expression by virtue of a direct inhibition of DNA
topoisomerase I [12].
Studies revealed that OAS is not only an ATP polymerase and should be defined as 2′-nucleotidyl
transferase [13–15]. In the experiments with partially purified OAS preparations isolated from
different sources (mammalian or chicken), the enzyme was able to catalyze in vitro transfer of the
5′-nucleotidyl moiety of a donor nucleoside triphosphate NTP (N is G, U, C, T, dA, dG, dC, dT) to a
2′-OH end of an acceptor molecule, such as ATP, NpA (3′,5′), NAD+ or tRNA. The addition of a donor
nucleoside triphosphate other than ATP terminates the chain elongation. The requirement for the
acceptor site was defined as an AMP group linked in a RpA configuration, where R stands for
pyrophosphate, NAD+, oligomeric or polymeric primers [15]. By using purified preparations of human
OASs (69-kDa and 100-kDa isoforms), Marie et al. showed the capacity of the human enzyme to use
GTP as an alternative substrate for OAS, as donor, and also as acceptor molecules [16]. However, GTP
was a poor substrate compared with ATP; in the presence of ATP, guanylate moiety was transferred as
donor on a 2-5A matrix followed by the release of the product [16]. Probably the best known natural
2′,5′-phosphodiester bond (between adenylate and guanylate moieties) is the one that forms in the
lariat RNA intermediate during gene splicing [17]; as OAS can also be located in the nucleus, the role
of OAS in generating that bond has been suggested [18,19]. Sperling et al. proposed that OAS activity
might be involved in pre-mRNA splicing [18].
Despite the elucidated nucleotidyltransferase properties of OAS, the natural occurrence of
2′,5′-oligonucleotides other than 2-5A has never been demonstrated. Furthermore, the biological role
of 2′,5′-linked heteronucleotides, which might be synthesized in cells, is not known [1,15,20,21].
Based on their structural motifs OAS proteins belong to the ancient group of nucleotidyl-
transferases, DNA polymerase β-like nucleotidyltransferase superfamily [22–24], being unique in
catalyzing the formation of 2′,5′-phosphodiester linkage instead of the usual 3′,5′-linkage. Studies of
OAS gene distribution among metazoans have shown their wide occurrence [25,26]; however, the
genomes of several organisms, even of those belonging to certain vertebrate lineages (e.g., some

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teleost fishes and the amphibian Xenopus tropicalis) do not reveal their presence. The OAS genes have
evidently been lost in some branches of the evolutionary tree of life [21,26–28].
Kuusksalu et al. have demonstrated the presence of OAS in the evolutionarily most basal
multicellular animals, sponges [29,30]. Several cDNAs encoding oligoadenylate synthetases have been
cloned from sponges [31–34]. Besides, a recombinant enzyme from Geodia cydonium was
characterized for its OAS activity [35]. The constructed phylogenetic tree reveals that the known
sponge OAS sequences fall into the same branch; vertebrate OASs are more distantly related [34].
Considering that the homology between primary structures of OAS from vertebrates and sponges is
rather low and their respective genomic structures are different, sponge OASs may be classified as a
distinct subgroup of OAS [36].
The role of OAS in sponges is still not known. The participation of this enzyme in responses to
environmental stressors [37] or in host-defense reactions (including bacterial and viral infection) has
been suggested [32,34]. However, the exact signalling pathways leading to any of these responses have
not been elucidated. The genes that define the interferon response in mammals (including those for
IFNs themselves) are absent from the known invertebrate genomes [38].
OAS activity can be detected in a variety of marine sponges (Demospongia), which exhibit different
2–5A synthesizing capacities and product profiles [39]. Compared with the enzymes of the mammalian
OAS family, OASs from some sponge species (G. cydonium, Chondrosia reniformis, Chondrilla
nucula) exhibit a surprisingly high activity [39]. The specific activity of OAS from these sponge crude
extracts is comparable or greater than that of the IFN-stimulated human (HeLa) or murine (L929) cells
[37,40]. In addition, the natural occurrence of 2-5A, in the amounts that were characteristic of
IFN-stimulated virus-infected cells, has been found in G. cydonium [29,37].
This study shows that two other marine sponges, Thenea muricata and Chondrilla nucula, can
accumulate 2-5A in high amounts. We demonstrate that besides 2-5A these sponges are capable of
synthesizing in vivo a series of 2′,5′-linked heteronucleotides, in which all four ribonucleotides (ATP,
GTP, UTP, CTP) are included. After dephosphorylation, they appeared in the form of A2′p5′N. NAD+
could be also used as in vivo substrate for OAS, since 2′-adenylated derivative of NAD+ was found in
the extract of T. muricata. In addition to ATP and NAD+, GTP was used in vivo as an acceptor
molecule for oligomerization to produce G2′p5′N (N is A or U). Reactions performed under in vitro
conditions with these substrates confirmed the results obtained in vivo.
Interestingly, our present study of the substrate specificity revealed that the sponge OAS is also
able to catalyze (2′-5′) oligomerization of pyrimidine nucleotides. Therefore, as a result of nucleotidyl-
transferase activity of the sponge OAS, which is considerably different from that of vertebrate
enzymes, a variety of 2′,5′-linked oligonucleotides can be obtained under in vitro conditions.
In the light of the new data obtained in the study we suggest that closer studies of OASs from
sponges, the most basal multicellular animals, would help us to elucidate original function(s) of the
enzymes of this nucleotidyltransferase family. On the other hand, such data could be useful for
understanding the functions of mammalian OASs that remain outside the scope of RNase L.

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2. Results and Discussion
2.1. Analysis of sponge perchloric acid extracts
The HPLC method for analyzing cellular nucleotides after their extraction from animal cells or
tissues is widely used for quantification of adenine nucleotides. In the course of such a study for which
a series of samples of the sponge T. muricata were used we found that this sponge contains
considerable amounts of diadenylates with 2′,5′-phosphodiester linkage. These diadenylates were
present in the perchloric acid extract in various 5′-phosphorylated (tri-, di-, mono-) and
unphosphorylated ("core") forms, the proportions of which depended on the particular sample (Figure
1). In some samples minor amounts of 2′,5′-triadenylates were also present (Figure 2). Such data
indicate that the oligoadenylates, initially synthesized as 5′-triphosphate derivatives resulting from
OAS activity exhibited by the sponge were afterwards dephosphorylated to a different degree.
However, the mechanisms that regulate phosphatase activities in particular sponges are not known yet.
Figure 1. The occurrence of 2′,5′-diadenylates in the form of tri-, di- and monophosphates,
and "core" forms in perchloric acid extracts of T. muricata.
HPLC analysis was performed by applying gradient 1 (see Experimental section). HPLC
chromatograms correspond to approximately 6 mg of the sponge wet weight. Blue, red and green
colored lines represent samples which were cultivated in laboratory tanks for different periods
of time.
Treatment of the extract with alkaline phosphatase simplified the subsequent analysis as all
compounds were separated in their "core" forms (Figure 2). Quantification of adenylates revealed that
their amounts varied by an order of magnitude between different samples of T. muricata (see
Experimental section, 3.1): the amount of 2′,5′-linked diadenylates (A2′p5′A) was 2–50 pmol/mg wet
weight, whereas that of adenosine (reflecting the total amount of adenine nucleotides ATP, ADP and
AMP) varied from 2 to 80 pmol/mg wet weight.

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Figure 2. The occurrence of 2′,5′-linked nucleotides in a perchloric acid extract of T. muricata.
HPLC analysis was performed by applying gradient 2 (see Experimental section). HPLC
chromatogram corresponds to approximately 8 mg of the sponge wet weight. Blue line represents
the nucleotide profile of a perchloric acid extract of the sample, red line–the product profile of the
same sample after dephosphorylation of the extract. Inset: the analysis of the sample by applying
HPLC gradient 1. The peaks P1-P3 were collected and analyzed: P1 involves A2′p5′G,
P2–NAD2′p5′A, P3–A2′p5′U and A2′p5′C.
Surprisingly, in many samples 2-5A was present in quantities that considerably exceeded those of
the total adenine nucleotide pool (Figure 2). Such a variation between the amounts of adenylates
obviously shows differences in the physiological status of sponges resulting i) from cultivation effects
and ii) from variability between different specimens. In accordance with the data published previously
[37], elevated levels of 2-5A evidently correspond to stress conditions for the sponge (data not shown).
A peak with retention time close to adenosine and UV spectrum (λmax = 256 nm) different from
oligoadenylates regularly accompanied high levels of A2′p5′A. Its intensity increased after
dephosphorylation of the extract and we presupposed the presence of some hetero-oligomer, which
might also have a 2′,5′-linkage. Therefore, the chromatographic fraction containing the unknown
compound (peak 1, Figure 2) was collected and subjected to respective chemical and enzymatic
treatments (see Figure 3) to establish the structure of the compound. Since alkaline hydrolysis of this
compound produced equimolar amounts of the products [(3′-AMP + 2′-AMP) and guanosine)], and
digestion with snake venom phosphodiesterase (PDE) produced equimolar amounts of 5′-GMP and
adenosine, the unknown compound has to be a (A, G) dimer. Resistance to the treatment with
ribonuclease T2 indicated that the dimer has a 2′,5′-internucleotide linkage instead of the usual
3′,5′-linkage.

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Figure 3. Determination of the primary structure and proof of 2′,5′-linkage in an oligoribonucleotide.
Alkaline hydrolysis of an oligomeric nucleoside produces 2′- and 3′-monophosphates while the
3′-terminal base is found as a nucleoside. Digestion with snake venom phosphodiesterase produces
5’-monophosphates while the 5′-terminal base is found as a nucleoside. Ribonuclease T2
hydrolyzes any 3′,5′-internucleotide linkage between ribonucleotides, but oligomers with
2′,5′-internucleotide linkage are resistant to this nuclease.
As a result of these treatments, the unknown compound was identified as adenylyl(2′-5′)guanosine
(A2′p5′G) (Figure 4, 1). Subsequent mass-spectrometric analysis (m/z 611.2) confirmed the identity of
the product.
Figure 4. Structures of compounds 1–6.
The mean content of A2′p5′G in different samples was about a magnitude lower than that of
A2′p5′A. A positive correlation observed between the amounts of A2′p5′A and A2′p5′G in different
samples of T. muricata (Figure 5) indicated that formation of the (A, G) dimer was directly related to
the activity of OAS in a particular sponge sample. Such a finding inspired us to look more carefully at
minor components present in different samples in order to search for additional oligonucleotides with
2′,5′-linkage.

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Figure 5. A correlation between the amounts of A2′p5′A and A2′p5′G in different samples
241
(n = 64) of T. muricata.
Dephosphorylated perchloric acid extracts of T. muricata were analyzed by HPLC. The amounts of
2′,5′-linked compounds were calculated from the same HPLC runs.
The area of a peak next to NAD+ (peak 2, Figure 2) correlated positively with the A2’p5’A peak as
well (not shown), but differently from A2′p5′A, A2′p5′G and NAD+ peak, its area did not increase by
phosphatase treatment of the extract. The UV spectrum of that peak was very similar to NAD+. Such a
behavior (including the observed retention time) would be expected for Nir-5′pp5′-Ado-2′p5′-Ado
(NAD2′p5′A), a compound formed from NAD+ and ATP by mammalian OAS [15]. MALDI-MS
analysis of this compound in positive ion mode was in agreement with NADpA (m/z 993.1; 992 Da for
neutral molecule). However, since NAD+ molecule has two ribose moieties with free 3′ and 2′ OH
groups, 5′-AMP (from the substrate ATP) might bind, alternatively, to nicotinamide ribose moiety.
The negative ion mode MALDI-MS analysis proved to be useful in solving this problem. The analysis
of NAD+ in negative ion mode yielded the strongest signals with m/z 540 (the ionized molecule
lacking nicotinamide) and 426 (corresponds to the fragment of ADP), whereas the peak of expected
mass (m/z 662) was low. Appearance of NAD+ in negative ion mode MALDI-MS analysis as
fragments with m/z 540 > 426 has been reported, while the signal with m/z 662 has not been detected
at all [41]. Similarly to NAD+, NADpA in negative ion mode yielded the strongest signals with m/z
869.1 and 755.1, together with the low intensity signal that corresponded to the expected mass with
m/z 991.1. On the basis of the fragments formed in both cases we can conclude that the adenylate
additional to NAD+ in NADpA was bound to adenine nucleotide moiety of the NAD+ molecule. Since
NADpA isolated from the sponge extract was resistant to RNase T2 treatment, the identified
compound was Nir-5′pp5′-Ado-2′p5′-Ado (Figure 4, 2).
NAD+ has been reported to be a preferred acceptor molecule for the OAS in human (HeLa) cell
extracts, however, the respective 2′-adenylated derivatives of NAD+ were not detected in the extracts
synthesizing 2-5A [42].
Another small peak in the samples with higher A2′p5′A content was found whose area increased
after treatment of the extract with alkaline phosphatase (peak 3, Figure 2, inset). As a result of the
respective treatments (Figure 3) and mass-spectrometric analysis, two compounds in the collected
chromatographic fraction were identified: one of them was adenylyl(2′-5′)uridine (A2′p5′U)

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(Figures 4, 3) (m/z 572.1) and the other was adenylyl(2′-5′)cytidine (Figure 4, 4) (A2′p5′C)
(m/z 571.1) (Figure 4). From these two compounds A2′p5′U was prevalent, its molar amount was
about two orders of magnitudes lower than that of A2′p5′A.
Thus, dimers containing all four ribonucleotides in the form of A2′p5′N (N is A, G, U or C) were
found in dephosphorylated extracts of T. muricata, the molar content of these compounds decreasing
in the order: A2′p5′A > A2′p5′G > A2′p5′U > A2′p5′C. Although the exact meaning of the synthesis of
such 2′,5′-linked oligomers for the sponge physiology is not known, decrease in cellular nucleotide
content, especially that of ATP, would have negative impact on total metabolic activity and could
indicate stress conditions for the sponge.
To expand the knowledge about the natural occurrence of 2′,5′-linked heteronucleotides in sponges,
the detectable levels of these metabolites could be found, guided by the example of T. muricata, in
sponges with high 2-5A content. The occurrence of 2-5A has been shown only for G. cydonium, using
radioimmunoassay method for establishing its concentration [29]. The content of 2-5A in G. cydonium
that was determined in the present study was up to 5 pmol per mg wet weight, being in the same range
with values found earlier. Two other sponges that had previously demonstrated high OAS activities
[39] were studied for their 2-5A content. In C. reniformis, the amount of 2-5A was about 2 pmol per
mg wet weight, whereas in C. nucula 2-5A was accumulated in higher amounts (14 and 3 pmol per mg
wet weight for dimer and trimer, respectively) (Figure 6). Therefore, C. nucula was chosen for
comparative studies of the occurrence of 2′,5′-linked heteronucleotides.
The presence of A2′p5′G (peak 1, Figure 6) was easily detected in dephosphorylated extracts of C.
nucula and also confirmed as described above. Similarly to T. muricata, the amount of A2′p5′G (which
also increased by phosphatase treatment) in the extract of C. nucula was about a magnitude lower than
that of A2′p5′A.
To establish other possible 2′,5′-linked oligomers in the extract of C. nucula, chromatographic
fractions (18 fractions, á 0.5 mL) in the region between guanosine and (2′-5′)ApApA peaks (Figure 6)
were collected and analyzed. The following hetero-dimers (migrating together with some other
components of the fraction) were identified in the analyzed region: peak 3 (23.5–24 min)–A2′p5′U
(Figure 4, 3) (m/z 572.1) and A2′p5′C (Figure 4, 4) (m/z 571.1), together with traces of adenosine and
A2′p5′A; peak 4 (17.5–18min)–guanylyl(2′-5′)uridine (G2′p5′U) (Figure 4, 5) (m/z 588.1) as a minor
component migrating together with dGuo; peak 5 (19–19.5 min)–guanylyl(2′-5′)adenosine (G2′p5′A)
(Figure 4, 6) (m/z 611.1) as a minor component migrating together with an unknown compound
(UVmax 261 nm). The molar content of 2′,5′-linked dimers in the extract that was calculated by peak
areas and from the data of alkaline hydrolysis of the collected fractions decreased in the order:
A2′p5′A > A2′p5′U > A2′p5′G > A2′p5′C > G2′p5′U ≥ G2′p5′A.
In comparison with T. muricata, two additional compounds with the formula G2′p5′N (N is A or U)
were found in the extract of C. nucula. In order to synthesize such hetero-oligomers, GTP has to be
bound at acceptor site and ATP at donor site of the OAS. Since ATP is a prevalent ribonucleotide in
cells (see also Figure 6 for the content of respective nucleosides), the affinity of GTP towards OAS
from C. nucula should be considerable.
To date, no data are available in the literature about the ability of a sponge OAS to catalyze the
formation of co-oligomers of ATP and another ribonucleotide. Therefore, to verify the hypothesis of in

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vivo synthesis of the 2′,5′-heteronucleotides in sponges, in vitro reactions were performed with NAD+
and four ribonucleotides, used alone or in mixtures with ATP.
Figure 6. The occurrence of 2′,5′-linked compounds in a perchloric acid extract of C. nucula.
HPLC analysis of a dephosphorylated sponge extract was performed by applying gradient 2 (see
Experimental section). HPLC chromatogram corresponds to approximately 30 mg of the sponge
wet weight. The peaks P1, P3, P4 and P5 were collected and analyzed: P1 involves A2′p5′G,
P3-A2′p5′U and A2′p5′C, P4–G2′p5′U and P5–G2′p5′A.
2.2. The formation of 2′,5′-linked oligomers in vitro
The extracts with specific activities of 138 pmol ATP/min per µg protein and 1.64 nmol ATP/min
per µg protein were used in the assays for T. muricata and C. nucula, respectively. NAD+ alone
remained intact in the extract, however, other ribonucleotides which were used alone as substrates
instead of ATP were easily converted to their respective 2′,5′-linked oligomers. In the case of GTP, the
specific activity was 43 pmol/min per µg protein for T. muricata and 2.34 nmol/min per µg protein for
C. nucula. Unexpectedly, pyrimidine nucleotides UTP or CTP were utilized quite efficiently to
produce the respective 2′,5′-linked oligomers. The calculated specific activity for T. muricata was
54 pmol UTP/min per µg protein or 38 pmol CTP/min per µg protein. For C. nucula, the specific
activities relative to pyrimidine nucleotides were 69 pmol UTP/min per µg protein and 13 pmol
CTP/min per µg protein.
These data show that the specific activity of the ribonucleotides towards OAS decreases in the
order: ATP > GTP ≅ UTP ≅ CTP or GTP ≅ ATP >> UTP > CTP for T. muricata and C. nucula,
respectively.
The product profiles formed from the ribonucleotides were studied under the assay conditions
where most of the given substrate was utilized for oligomerization. In these conditions, 2′,5′-linked
oligomers up to pentamers (ATP), tetramers (GTP) and trimers (UTP and CTP) were formed (Table 1).
The relative amounts of the formed oligomers are presented in Table 2. As it can be seen, purine
nucleotides were better substrates for chain elongation than pyrimidine nucleotides. The quantitative
data of oligomerization of CTP by the C. nucula extract are not included in Table 2 due to a
deaminating activity which was present in the extract. This resulted in a quite extensive conversion of
CTP to UTP (with a specific activity of 22.6 pmol/min per µg protein) and in the production of a
mixture of 2′,5′-linked products consisting of C and U homo-oligomers as well as of (C, U) hetero-
oligomers.

Mar. Drugs 2010, 8
Table 1. Analytical forms of 2′,5′-linked homo-oligoribonucleotides.
244
Substrate
(2′-5′) Oligomer*1
ApA
Retention time, min*2
m/zexp*3
595.1
924.1
1253.1
1582.3
627.1
972.1
1317.3
549.0
855.1
547.0
852.0
ATP
22.2
22.5
21.9
21.0
14.9
17.5
17.9
15.1
16.3
10.6
11.4
ApApA
ApApApA
ApApApApA
GpG
GpGpG
GpGpGpG
UpU
GTP
UTP
CTP
UpUpU
CpC
CpCpC
*1 the products were analyzed in their dephosphorylated forms.
*2 retention times correspond to the applied gradient 1 (see Experimental Section).
*3 experimentally obtained mass signal in negative ion mode MALDI-MS.
Table 2. The relative molar amounts of 2′,5′-linked homo-oligomers (%) formed from
different ribonucleotides.
Thenea muricata
Substrate
Incubation
Monomer*
Dimer
Trimer Tetramer Pentamer
time, h
17
ATP
GTP
UTP
CTP
3.4
5.7
5.1
7.4
27.6
67
85.6
88.1
48.7
24.1
9.3
18.5
3.3
-
-
1.9
20
20
20
-
-
-
4.5
Chondrilla nucula
ATP
GTP
UTP
1
1
18
4.1
10.1
13.6
31.3
50.6
76.4
59.6
35.7
10
5
3.6
-
-
-
-
*–the amount of the respective mono-nucleotide remaining at the end of incubation.
Thus, as illustrated by the example of T. muricata and C. nucula, the substrate specificity of OAS
depends on the particular sponge species. The inter-species comparison of known sponge OAS protein
sequences refers to a very low similarity between these proteins, most probably due to their
independent evolution after the divergence from their common ancestor [36]. Although the OAS genes
from T. muricata and C. nucula have not been sequenced, different affinities of ribonucleotides
towards OAS might be explained by differences in the enzyme structures of the respective sponges.
To study the formation of hetero-oligomers, ATP was combined with GTP, UTP, CTP or NAD+ in
roughly equimolar concentrations. Since G2′p5′U was identified after phosphatase treatment of
perchloric acid extract of C. nucula, the combination of GTP and UTP was used additionally in case of
this sponge. Taking into account the relative ability of a sponge OAS to catalyze the oligomerization
of these substrates when used alone (see above), the nature and multiplicity of the products formed by

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one or another of the sponges in the experiments with the mentioned mixtures is not surprising
(Table 3).
Table 3. 2′,5′-linked heteronucleotides formed in vitro from the substrates added to the
extract of the sponge Thenea muricata (1) or Chondrilla nucula (2).
3
Substrates
(2′-5′) Oligomer*1
GpGpA
GpGpApA
GpA
Retention time, min*2
m/zexp
*
Sponge
ATP + GTP
16.7
16.9
17.5
18.1
19.2
19.4
20.2
20.3
20.7
17.7
18.4
18.8
19.1
19.9
19.9
20.9
13.7
15.5
15.5
19.5
19.9
19.9
20.9
21.0
14.6
15.7
16.0
13.2
956.1
1285.2
611.1
940.1
956.1
611.1
940.2
956.2
940.2
878.0
1230.1
572.0
901.0
901.0
1230.1
572.0
876.1
571.1
900.1
1229.2
901.1
900.1
572.1
571.1
933.1
1278.1
588.1
993.1*5
2
2
1, 2
1, 2
2
GpApA
ApGpG
ApG
1, 2
1, 2
2
ApApG
GpApG
ApGpA
UpApU
UpApApA
UpA
2
ATP + UTP
1
1
1
UpApA
ApApU
ApApApU
ApU
1
1, 2
1
1, 2
1
ATP + CTP
CpApC
CpA
1
CpApA
ApApApC
ApApU*4
ApApC
ApU*4
1
1
2
1, 2
2
ApC
1, 2
2
GTP + UTP
GpGpU
GpGpGpU
GpU
2
2
ATP + NAD+
NADpA
1, 2
*1, **2, ***3–see explanations of Table 1.
*4–these products were formed due to a deaminating activity of the extract of C. nucula which
resulted in a partial conversion of CTP to UTP (see above).
*5–mass signal in positive ion mode MALDI-MS.
The results demonstrate that adenylate was preferred in 5′-terminal position in hetero-oligomers,
and the yield of dimeric product A2′p5′N was generally the highest (Figure 7A, C, D). As an
exception, there was only a slight preference for ATP in 5’-terminal position when the ATP + GTP
mixture was added to the C. nucula extract (Figure 7 B). In the reaction with GTP + UTP, GTP was an
efficient acceptor molecule while UTP terminated the chain (Table 3). In the case of T. muricata,

Mar. Drugs 2010, 8
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where the difference between OAS specific activities towards purine and pyrimidine nucleotides was
smaller than that observed for C. nucula, pyrimidine nucleotides could also serve (though with a
smaller efficiency) as acceptor molecules for oligomerization of ATP in the presence of UTP or CTP
(Table 3, Figure 7C).
Figure 7. The oligomerization of ATP in the presence of GTP (A, B) or UTP (C, D) by
OAS from the extract of T. muricata (A, C) or C. nucula (B, D) under in vitro conditions.
Substrates were used in the following concentrations: 0.86 mM ATP, 0.67 mM GTP, 0.76 mM
UTP. Incubation time was 1 h for C. nucula and 17 h for T. muricata. HPLC analysis of
dephosphorylated reaction mixtures was performed by applying gradient 1 (see Experimental
section). The chromatographic peaks were collected and analyzed. The numbered compounds in
panels are: A–1 (G2′p5′G2′p5′G + G2′p5′A2′p5′A), 2 (A2′p5′A2′p5′G), 3 (A2′p5′A2′p5′A);
B–1 (G2′p5′G2′p5′A), 2 (G2′p5′G2′p5′A2′p5′A), 3 (G2′p5′G2′p5′G), 4 (G2′p5′A2′p5′A),
5 (A2′p5′G2′p5′G), 6 (A2′p5′A2′p5′G), 7 (G2′p5′A2′p5′G), 8 (A2′p5′G2′p5′A), 9 (A2′p5′A2′p5′A);
C–1 (m/z 1207.1, the sequence of nucleotides not analyzed),
2 (U2’p5’A2’p5’U),
3 (U2′p5′A2′p5′A2′p5′A), 4 (U2′p5′A2′p5′A), 5 (A2′p5′A2′p5′U + A2′p5′A2′p5′A2′p5′U),
6 (A2′p5′A2′p5′A2′p5′A + A2′p5′A); D–1 (A2′p5′A2′p5′U), 2 (A2′p5′A).
Thus, these data provide a clear proof for the ability of OAS to catalyze in vivo the production of
2′,5′-heteronucleotides. The results show that besides ATP, other nucleotides are efficiently used for
the formation of 2′,5′-linked oligomers under in vitro conditions. Especially, the usage of pyrimidine
nucleotides as acceptor molecules for the oligomerization refers to the substrate specificity of the
sponge OAS that is remarkably different from that of vertebrate OASs.
It has previously been proposed that OAS proteins have evolved from a common ancestor of
archaeal CCA-adding enzymes and poly(A) polymerases [24,25]. The OAS proteins show the highest
similarity with the class I CCA-adding enzymes, which not only catalyze the synthesis of the CCA-
sequence to the 3′-tRNA end, but are also capable of poly(C) synthesis [43]. Interestingly, OASs from
T. muricata and C. nucula are capable of catalyzing the synthesis of 2′,5′-linked homo-oligomers not
only from ATP, but also from GTP, UTP and CTP. These findings indicate that OASs from sponges
have broader nucleotide specificity than that known for vertebrate OASs. Further studies of these
enzymes from sponges allow for better characterization of their nucleotidyltransferase properties.

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Moreover, research in this field may help to understand the function of OAS proteins in sponges,
the most basal multicellular animals. Taking into consideration that several OAS genes are expressed
in a sponge and that, on the other hand, OAS protein sequences in different sponge species differ
considerably from one another [36], the functions of OASs in sponges may also be diversified. Though
at present suggestions about the role(s) of OASs in sponges are speculative, one can consider a
possible role of these enzymes in the control of cell growth, which may be regulated by different signal
pathways depending on the inducing factor. Another intriguing aspect is the participation of OASs in
immune responses of sponges against bacteria or viruses as also proposed earlier [32,34,36]. The
elucidation of the pathways involved in these host-defence reactions would give us valuable
information about the innate immune system of invertebrates. In its turn, thorough studies of OASs
from sponges could enlighten some aspects of the suggested multifunctionality of OAS proteins in
vertebrates.
3. Experimental Section
3.1. Sponges
The marine sponges Thenea muricata (Demospongiae) were collected on a soft bottom sediment off
Korsfjord (close to Bergen, Norway) at 300 m depths (60º 07’ 44 N, 004º 49’ 9 E). The sponges were
either fixed directly in liquid nitrogen or maintained in tanks with running sea water for establishing a
laboratory-based culture and then fixed. The total number of analyzed samples (corresponding to
sponge specimens) was 64. The samples were kindly provided by Hans Tore Rapp from Bergen
University. The marine sponges Chondrilla nucula, Geodia cydonium and Chondrosia reniformis
(Demospongiae) collected from the Tyrrhenian Sea (Napoli Bay) were kindly provided by Dr. S. de
Rosa (CNR, Napoli, Italy). The specimens of G. cydonium collected from the Adriatic Sea (Rovinj,
Croatia) and C. reniformis collected from Cala Montgo, L’Escala (Spain) were gifts from Prof. Dr. W.
E. G. Müller (University of Mainz, Germany) and Dr. M.-L. Schläppy (MPI for Marine Microbiology,
Bremen, Germany), respectively. The animals were cut into pieces, frozen in liquid nitrogen and kept
at -70 °C until their further use.
3.2. Sample homogenization
The sponge tissue was disrupted by means of TissueLyser (Qiagen, Retsch GmbH, Germany).
Grinding jars were pre-chilled in liquid nitrogen and the frozen samples were pulverized in a high-
speed regime (at 30 Hz) for 1 min.
3.3. Preparation of extracts
3.3.1. Perchloric acid extract
0.5 mL of an ice-cold 0.6 M HClO4 solution was added per about 100–150 mg of a powdered
frozen sample and mixed vigorously. The extract was additionally vortexed and kept on ice for 10 min.

Mar. Drugs 2010, 8
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After centrifugation for 5 min (10,000 g, 4 °C), the supernatant was removed and neutralized with
5 N KOH to about pH 7. The KClO4 precipitate formed was removed by centrifugation (5 min,
10,000 g, 4 °C), the supernatant was collected and submitted to HPLC analysis either directly or after
treatment with shrimp alkaline phosphatase (SAP, see 3.6). In order to identify minor components
present in the extracts, the extract was first concentrated (10–30 fold) by lyophilization under reduced
pressure (SpeedVac Concentrator, Savant) and then analyzed, either directly or after treatment with
SAP, by the HPLC method.
3.3.2. Protein extract
The powdered frozen sponge tissue was extracted for 15 min at 0 °C with 10 mM HEPES (pH 7.5)
buffer containing 100 mM KCl. The insoluble pellet was removed by centrifugation (5 min, 10,000 g,
4 °C) and the supernatant was used for activity assays. The protein concentration of the extract was
measured by a modified Bradford method [44]. The extracts with protein concentrations of 2.5 mg/mL
and 2.8 mg/mL were used for T. muricata and C. nucula, respectively.
3.4. 2′,5′-Oligoadenylate synthetase activity assays
The reaction mixtures containing 20 mM Tris-HCl, pH 8, 50 mM KCl, 5 mM MgCl2, the sponge
protein extract (final protein concentration from 0.005 to 0.1 mg/mL) and the substrate(s) (ATP, GTP,
UTP, CTP, NAD+) were incubated at 37 °C for required periods of time (5 min–20 h). The reaction
was stopped by heating (4 min, 95 °C). Substrates were used in the following concentrations: 0.86 mM
ATP, 0.67 mM GTP, 0.76 mM UTP, 0.78 mM CTP and 0.97 mM NAD+.
For the determination of specific activity where the appearance of the firstly synthesized dimer was
registered, short reaction times (5–15 min) and low sponge protein concentrations (0.05 to 0.005 mg
protein/mL) were suitable. To study the total pool of oligomeric products formed in different reactions,
the protein concentrations were 0.1 mg/mL and incubation times were longer. Overnight incubations
(17–20 h) for all substrates were suitable for T. muricata. For C. nucula, 1 h incubations were used
with all substrates or their combinations, except for pyrimidine nucleotides UTP or CTP when used
alone. In the latter case, the incubation time was 18 h.
Products of the reactions were analyzed either directly or after dephosphorylation of the reaction
mixture (see 3.6).
3.5. HPLC analysis
The analysis was performed with an Agilent 1100 HPLC instrument (Agilent Technologies, USA)
on a C18 reverse-phase column (Phenomenex Luna C18(2), 250 × 4.6 mm, 5 μm; pre-colum
SecurityGuard C18, 4 × 3 mm). Eluent A was 50 mM ammonium phosphate pH 7.0 and eluent B was
50% methanol in water. The products were separated at 30 °C in a linear gradient of eluent B (0–40%,
20 min); the column was equilibriated with eluent A before the next injection (10 min) (gradient 1).
Alternatively, the column was thermostated at 20 °C and the linear gradient of eluent B (0–60%,
30 min) was used (gradient 2). The absorption was measured at 260 nm. The adenylates and other

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products were identified by comparing their retention times with those of authentic compounds and by
their UV spectra (using previously created UV spectra library). The retention times of the respective
compounds with 2′,5′- and 3’,5’-linkage differed substantially, the latter having longer retention in the
column. For example, by applying gradient 1 the retention time of G2′p5′U was 16 min, whereas that
of G3′p5′U (Sigma) was 20 min; the retention time of A2′p5′U was 20.9 min versus 23 min for
A3′p5′U (Sigma).
The products were quantified by integrating the corresponding peak areas (software of Agilent
Technologies). Taking into account the molar absorption coefficients of the compounds at 260 nm,
molar concentrations were calculated. Molar absorption coefficients for 2′,5′-adenylates have been
given previously [45]. The following molar absorption coefficients were used for other oligomers:
G2′p5′G–2.11 × 104, G2′p5′G2′p5′G–3.16 × 104, G2′p5′G2′p5′G2′p5′G–4.21 × 104, U2′p5′U–1.8 × 104,
U2′p5′U2′p5′U–2.7 × 104, C2′p5′C–1.36 × 104, C2′p5′C2′p5′C–2.04 × 104.
3.6. Analysis of the composition of oligoribonucleotides by means of chemical and enzymatic
hydrolysis
The composition of oligoribonucleotides was determined by analysis of the core oligomers after
removal of the 5′-phosphate groups by alkaline phosphatase treatment. The dephosphorylation reaction
was performed by adding the appropriate amounts of 10 x reaction buffer (100 mM MgCl2, 200 mM
Tris-HCl, pH 8) and shrimp alkaline phosphatase (SAP, USB Corporation) at final concentration of
0.02–0.04 U/µL and incubating the mixture at 37 °C for 1 h; thereafter the enzyme was inactivated
(5 min, 95 °C).
3.6.1. Alkaline hydrolysis
Oligomer(s) of the collected chromatographic fractions were treated for 10 min at 95 °C with 0.3 M
NaOH, placed on ice and neutralized to about pH 7 by adding the appropriate amount of 2 N HCl. The
products of hydrolysis were identified by the HPLC method. The length of an oligomer was evaluated
by means of quantification of the resulting products of hydrolysis.
3.6.2. Snake venom phosphodiesterase treatment
Oligomers were treated in 50 mM Tris-buffer, pH 9 for 1 h at 37 °C with snake venom
phosphodiesterase (phosphodiesterase from Vipera lebetina venom was kindly provided by Dr. J
Siigur, NICBP, Tallinn) at final concentration of 0.04 mg/mL. The enzyme was inactivated by heating
(5 min, 95 °C). Partial digestion of an oligomer by using shorter incubation times (5–30 min) and
lower enzyme concentrations (0.004–0.008 mg/mL) allowed us to determine the nucleotide
composition on the basis of resulting shorter fragments of the oligomer. The products were analyzed
by the HPLC method.

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3.6.3. Ribonuclease T2 treatment
The oligomers under investigation were incubated overnight in 50 mM sodium acetate buffer, pH 4,
at 35 °C with RNase T2 (Invitrogen) at final concentration of 0.01 U/µL. Reference
oligoribonucleosides A3′p5′A (Sigma), A3′p5′A3′p5′A (BioSpring, Germany), A3′p5′U (Sigma),
G3′p5′U (Sigma) were incubated for 1 h in the same conditions; all oligomers were totally degraded.
The products of hydrolysis were identified by the HPLC method.
3.7. MALDI-MS analysis
The collected HPLC fractions were directly subjected to mass-spectrometric analysis. The analysis
was carried out with a Bruker Daltonics Autoflex II TOF/TOF instrument in a reflector negative mode.
THAP (2,4,6-trihydroxyacetophenone) matrix solution was used as described previously [39].
[THAP-H+], [ApA-H+], [ApApA-H+] and [ApApApA-H+] with m/z of 167, 595.14, 924.1 and
1253.15, respectively, were used as external calibrants. DHB (2,5-dihydroxybenzoic acid) matrix
solution was used for registration of masses in the region with m/z from 547 to 549. The mass signals
of NAD+ and NAD2′p5′A were registered also in a reflector positive mode, using THAP as a matrix
solution. The discrepancies between registered and theoretical monoisotope masses were in the range
of 0.1 Da.
4. Conclusions
The perchloric acid extracts prepared from the marine sponges T. muricata and C. nucula reveal the
occurrence of 2′,5′-linked oligoadenylates, mostly in dimeric forms. As established by the example of
T. muricata, concomitantly with accumulation of 2′,5′-diadenylates, several 2′,5′-linked heterodimers
were also detected in the extracts. In particular, a positive correlation observed between the amounts of
adenylyl(2′-5′)adenosine and adenylyl(2′-5′)guanosine (or NAD2′p5′A) refers to the formation of
2′,5′-linked heterodimers due to the activity of OAS in a particular sponge sample. After
dephosphorylation of perchloric acid extracts, these 2′,5′-heterodimers were identified as A2′p5′N,
where N is G, U or C (C. nucula, T. muricata); as NAD2′p5′A (T. muricata) and as G2′p5′N, where N
is A or U (C. nucula). The case of G2′p5′U gives evidence for the presence of non-adenylate 2′,5′-
linked compound in sponge cells as a result of OAS activity. Thus, in addition to ATP and NAD+, GTP
was used in vivo as an acceptor for (2′-5′) oligomerization. Evidently, the formation of a particular
2′,5′-linked product by OAS depends on the characteristics of the enzyme and, on the other hand, on
bioavailability of its substrates in sponge cells.
The in vitro assays demonstrated that in T. muricata ATP was a preferred acceptor for the OAS,
while in C. nucula GTP was comparable with ATP as an acceptor for the enzyme. That can explain the
natural occurrence of 2′,5′-linked heteronucleotides in C. nucula, which were identified in the form of
G2′p5′A and G2′p5′U. Besides, the usage of pyrimidine nucleotides UTP or CTP as substrates in vitro
was quite unexpected, since (2′-5′) oligomerization of pyrimidine nucleotides by vertebrate OASs has
never been demonstrated. These results show that OASs from most basal multicellular animals,
sponges, have substrate specificity properties that are substantially different from those of vertebrate

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enzymes. With OASs from the marine sponges T. muricata and C. nucula, a variety of 2′,5′-hetero-
oligomers (up to tetramers) was obtained in vitro wherein all four ribonucleotides can serve as either
acceptor or as donor substrates for oligomerization. Notably, the substrate specificity of an OAS
depends on the particular sponge species, which might be explained by differences in the enzyme
structures of the respective sponges. Taking into account a great number of extant marine sponge
species, one could expect a much greater variability in the substrate specificity properties of sponge
OASs in general. Thus, a future finding of new naturally occurring 2′,5′-heteronucleotides in sponges
would be expected. On the other hand, the specific features of OASs from marine sponges open
perspectives for enzymatic synthesis of authentic samples of a variety of 2′,5′-linked heteronucleotides.
Acknowledgements
We wish to thank Hans Tore Rapp (University of Bergen, Norway) and Friederike Hoffmann
(Uniresearch AS, Bergen, Norway) for providing samples of Thenea muricata. Samples were collected
and processed in the framework of the joint project "Sponge risk assessment and monitoring-
SPONGRAM", funded by Statoil. This work was supported by the Estonian Ministry of Education and
Research (grant No. 0140108) and Estonian Science Foundation (grant No. 7421).
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