# Chaperone Activity of SARS-CoV-2 Nucleocapsid Protein: RNA Annealing and Destabilization Mechanisms

Loussine Zargarian, Xia Ai, Ze-Yu Song, Lara Perez-Gorgol, Esteban Couprie-Diaz, Xu-Guang Xi, Olivier Mauffret, Philippe Fossé

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## To cite this version:

Loussine Zargarian, Xia Ai, Ze-Yu Song, Lara Perez-Gorgol, Esteban Couprie-Diaz, et al., Chaperone Activity of SARS-CoV-2 Nucleocapsid Protein: RNA Annealing and Destabilization Mechanisms. Journal of Molecular Biology, 2025, 437 (19), pp.169312. 10.1016/j.jmb.2025.169312. hal-05370606

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# Chaperone Activity of SARS-CoV-2 Nucleocapsid Protein: RNA Annealing and Destabilization Mechanisms

Loussine Zargarian¹,†, Xia Ai²,†, Ze-Yu Song², Lara Perez-Gorgol¹,
Esteban Couprie-Diaz¹, Philippe Fossé¹, Xu-Guang Xi¹, and Olivier Mauffret¹,*

1 - Université Paris Saclay, Laboratoire de Biologie et Pharmacologie Appliquée, UMR 8113 CNRS, École Normale Supérieure de Paris Saclay, 91190 Gif sur Yvette, France

2 - College of Life Sciences, Northwest A&F University, Yangling, Shaanxi 712100, China

Correspondence to Olivier Mauffret: olivier.mauffret@ens-paris-saclay.fr (O. Mauffret)

https://doi.org/10.1016/j.jmb.2025.169312

Edited by Owen Pornillos

## Abstract

The recent global COVID-19 pandemic has prompted new research in coronaviruses. Despite the discovery of effective vaccines and therapeutic interventions, coronaviruses remain a threat to humans as the emergence of novel variants or new pathogenic coronaviruses is possible. The N protein or nucleocapsid protein, belonging to the virus' essential proteins, is mainly involved in the compaction and protection of the viral genome. Here, we show that the SARS-CoV-2 N protein is a very efficient chaperone protein of nucleic acids that aggregates nucleic acids and anneals well-folded and complementary oligonucleotides hairpins *in vitro*. Using fluorescence and gel shift electrophoresis methods, we showed the high ability of the protein to destabilize nucleic acid secondary structure and to anneal nucleic acid strands. This last activity needs the full-length protein as we demonstrate that protein fragments, while they could display some activities, are considerably less efficient. However, the ability of the full-length protein to destabilize small double-stranded RNAs is poor. Our results show that the N protein possesses a chaperone activity similar to the HIV-1 NCp7 and NCp9 nucleocapsid proteins known as powerful nucleic acid chaperons.

© 2025 CNRS. Published by Elsevier Ltd. This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).

Several years after the COVID-19 pandemic, the molecular mechanisms involving the SARS-CoV-2 proteins are incompletely known. The COVID 19 genome includes ORF1a, ORF1b and the remaining one-third of the genome has overlapping ORFs, encoding four major structural proteins, including S (spike glycoprotein), N (nucleocapsid protein), M (membrane protein) and E (envelope protein), and some accessory proteins -S-ORF3-E-M-ORF6-ORF7 (7a and 7b)-ORF8-ORF9b-N in order [1,2]. Several studies point to the essential role of mutations of N protein present in Delta and Omicron variants to increase viral particle infectivity [3,4].

The major role of this latter RNA-binding protein, known as the most abundant viral protein [5] is to

form a viral ribonucleoprotein complex (vRNP) permitting the compaction of the viral genome, its protection, its anchoring to the membrane of the viral particle [6–9]. Also, the packaging process could be related to the existence of a packaging signal localized in the viral genome and recognized by the N protein [7,10–13]. Additionally, this protein could be involved in replication and transcription processes [14–17]. More specifically, it is hypothesized that the N protein facilitates template switching required for the formation of sub-genomics RNA (sgRNAs) during transcription.

The SARS-CoV-2 N protein (419 a.a) is a highly dynamic protein with a modular organization of five different domains (Figure 1) [18–20]. Two of these domains are folded, the first one is the RNA
<figure><img src="image_4.png" /></figure>

<figure><img src="image_5.png" /><figcaption>**Figure 1. (A)** Predicted secondary structures for TAR (Transactivator response element) and cTAR derivatives used in this study. cTAR is the complementary sequence of TAR. For mini-cTAR-loop and mini-cTAR-stem, mutations relative to mini-cTAR are indicated in red. **(B)** Scheme of the N protein of SARS-CoV-2 used in this study; at the top of the figure, the five main domains of the protein are indicated (see the text for explanations) with a color code and indication of the amino acid numbering, below are indicated the four constructs that have been used in our study.</figcaption></figure>

binding domain (RBD, 50–174) [21–25], and the second, the dimerization domain (246–365), is responsible for dimerization of the protein [6,26–28]. In contrast, the three other domains are of a disordered nature, the N-terminal extension (NTE, 1–50) [5,20,29,30], the C-terminal extension (CTE, 365–419) [5,20] and finally the central linker (LINK, 174–246) [20,29,31] that connects the two folded domains. The LINK contains a serine/arginine (S/R) rich region [5,32–35] that harbors several phosphorylation sites. Several studies point to the fact that RBD is not the only domain interacting with RNA and that other domains also could also interact with RNA [18,27,29,30,36,37].

Recent studies on the N protein of SARS-CoV-2 and its biophysical properties showed that this protein drives liquid–liquid phase separation with RNA and forms RNA-dependent biomolecular condensates. Indeed, the modular nature of the protein, the presence of disordered domains and the ability of several parts of the protein to undertake protein–protein or protein-RNA interactions are properties typical of multivalent molecules known to favor liquid–liquid phase separation [5,13,20,28,38–42]. The formation of membrane-less biomolecular condensates is important for several viral processes such as viral replication, viral packaging and particle assembly [43,44].

The oligomerization status of the N protein is critical because several oligomeric forms have been identified: dimers, tetramers, octamers and higher oligomeric states [6,27,45–47]. The RNA binding could modify the degree of oligomerization of the protein in a dependent-RNA length phenomenon [47–49]. The dimerization process involves RBD [6,26,27], and several studies point to the role of a domain located in the linker as the tetramerization interface [45,50–52]. In addition,

the NTE and CTE domains appear to be able to modulate the oligomerization properties [27,53].

While many works have studied the RNA binding properties of N protein or its ability to form biomolecular condensates in the presence of RNA, very few studies have investigated its ability to reorganize RNA molecules. In a pioneering work on the N protein of transmissible gastroenteritis coronavirus (TGEV), Enjuanes et al. have shown that the protein behaves as a chaperone protein of nucleic acids able to anneal efficiently oligonucleotides [16,54]. These experiments suggest that the chaperone activity could be essential to mediate the TRS-cTRS (Transcription Regulating Sequence) interaction, a crucial step in the discontinuous synthesis of the subgenomic RNAs (sgRNAs) [15,16,55]. Furthermore, several other authors analyzed the ability of fragments of the N protein of SARS-CoV 2 [24,56] or Mouse hepatitis virus (MHV) [57,58] to destabilize RNA duplexes mimicking the TRS-cTRS duplex. In addition, a recent study demonstrates the involvement of the SARS-CoV-2 nucleocapsid protein in genome dimerization and the annealing of some parts of the genome with host cell miRNA [59].

In this study, we analyzed the ability of the SARS-CoV-2 nucleocapsid protein to hybridize self-folded oligonucleotides and to destabilize double-stranded stems. We showed, using two different methodologies, the excellent ability of the N protein to perform the annealing of folded and complementary structures with an efficiency similar to other viral proteins known as powerful nucleic acid chaperons. We also showed that the N protein aggregates the nucleic acids at relatively high concentrations. In contrast, the ability of the protein to destabilize double-stranded stems is not so efficient. We also found that selected domains
or portions of the protein are less efficient than the full-length protein in hybridizing complementary stem-loops. Altogether, our results indicate that N protein can be considered a powerful nucleic acid chaperon and we discuss the possible implications of this finding.

# Results

## EMSA assays highlights N1-419 as a potent facilitator of RNA-DNA annealing

To investigate whether the full-length SARS-CoV-2 nucleocapsid protein (named N1-419 in the study) can facilitate the annealing of nucleic acids, we used two hairpin oligonucleotide substrates (DNA and RNA) derived from the HIV-1 TAR sequence (Figure 1A) [60,61], as it has been done in other similar studies of viral nucleic acid chaperons (HIV-1, HBV) [61–65].

In a first assay to estimate the ability of N1-419 to facilitate base-pairing interactions between complementary nucleic acids, we used the mini-TAR RNA-DNA annealing assay (Figure 1A) using Electrophoretic Mobility Shift Assay (EMSA) [61]. In this assay, the heteroduplex product formed from the mini-TAR RNA and mini-c TAR DNA stem-loops is detected by native gel electrophoresis (Figure 2A). Without protein, the average yield of heteroduplex product was 5% (Figure 2B). NCP9, the precursor of the mature HIV-1 nucleocapsid protein (NCp7), was used as a positive control because it has strong annealing activity [66,67]. In the presence of increasing amount of N1-419, the rate of heteroduplex increased (Figure 2). The average yield of heteroduplex product was 73% at an N1-419 to nucleotide molar ratio of 1:7 (Figure 2B). Interestingly, the annealing activity of N1-419 was similar to that of NCP9.

Furthermore, we compared the activity of N31-210 and N240-419 proteins to promote the mini-TAR RNA-DNA annealing reaction (Figure 3). N31-210 contains the RBD domain and the part of the linker containing the Serine-Arginine (SR) region but not the leucine-helix (LH) region (Figure 1B). N240-419 contains the dimerization domain and CTE domains. As expected, at protein to nucleotide molar ratios greater than 1:28, the annealing rate was significantly greater with the N1-419 protein than with the N31-210 and N240-419 proteins (Figure 3B). Thus, at a protein to nucleotide molar ratio of 1:7, the annealing rate was 5.3-fold increased with N1-419 compared to the truncated proteins. At protein to nucleotide molar ratios greater than 1:28, the average yield of heteroduplex product was significantly increased with both truncated proteins compared to the control (Figure 3C). Therefore, both truncated proteins have slight annealing activity. However, their annealing activity was similar (Figure 3C).

## Fluorescence analysis reveals N1-419's efficient promotion of mini-TAR RNA-DNA annealing

In the following to investigate with additional details the process of mini-TAR – mini-c TAR annealing by N1-419, we used fluorescence methods relying on the FRET transfer (Forster Resonance Energy Transfer). Similarly to the EMSA experiments, we used two hairpin oligonucleotide substrates (DNA and RNA) derived from the HIV-1 TAR sequence [60,61], as shown in Figure 1A. The full-length N protein (N1-419) and its various fragments (Figure 1B) were assayed under identical experimental conditions. N1-260 fragment not used in the EMSA assays contains the entire linker domain.

The experiments were performed at 35 °C in conditions close to those for HIV-1 nucleocapsid proteins. In the first step, we quantified the Forster Resonance Energy Transfer (FRET) for two fluorophores, 5/6-carboxyfluorescein (F) and 5/6-carboxytetramethylrhodamine (TMR), located at the 3' and 5' extremities of the mini-c TAR DNA hairpin (Figure 1A), respectively. After the addition of the complementary mini-TAR RNA hairpin to the mini-c TAR DNA hairpin in the presence of a protein possessing a nucleic acid chaperone activity, the complementary hairpins anneal, and this reaction is typically associated with a decrease of FRET transfer between the two fluorescent probes [60,68,69]. The results are presented in Figure 4. The full-length protein (N1-419) and the N1-260 and N31-210 fragments at concentrations lower than one µM strongly decreased the FRET efficiency (EFRET). In contrast, the EFRET decrease was less significant with the N240-419 fragment, even at high protein concentrations. In addition, the concentration necessary to obtain the complete loss of the FRET was much lower for N1-419 (~180 nM) than for the N1-260 (~560 nM) and N31-210 (~450 nM) fragments. Note that the FRET level when no protein and RNA were present was close to 0.8, indicating a small opening of the hairpin extremity, expected to occur at 35 °C.

In another set of experiments, we analyzed the kinetic aspects of the reaction, a feature that is most easily inspected with fluorescence methods. The experiments were performed at various ratios of protein:DNA/RNA expressed in terms of protein:nucleotide molar ratio. A typical set of data is shown for a ratio of 1:29 (1 protein for 29 nucleotides) (Figure 5) and the comparison with kinetics of the annealing in absence of the protein is shown in Supplementary Figure S1. In these experiments, the DNA concentration was constant and RNA and protein concentrations increased accordingly to maintain a fixed protein-to-nucleotide ratio. At the three ratios tested (1:29, 1:15 and 1:8), the annealing process displayed two different phases, a fast one occurring during
<figure><img src="image_6.png" /><figcaption>**Figure 2.** Analysis of mini-TAR RNA-DNA annealing in the presence of NCp9 or N1-419. The annealing assays were performed as described in Materials and Methods. (A) The samples were analyzed by electrophoresis on 15% polyacrylamide gels at 20 °C in the TBE buffer. The 5'-end labeled mini-TAR RNA alone was incubated in the absence of protein (lanes C1) or in the presence of NCp9 (lane C3) or N1-419 (lane C4). The 5'-end labeled mini-TAR RNA was incubated with mini-cTAR DNA in the absence of protein (lanes C2) or in the presence of NCp9 (lanes 5-7) or N1-419 (lanes 11-13). The protein to nucleotide molar ratios were 1:28 (lanes 5 and 11), 1:14 (lanes 6 and 12) and 1:7 (lanes 4, 7, 10 and 13). Heat-denatured mini-TAR RNA (lane D) was used to identify the band corresponding to the monomeric form of mini-TAR. The monomeric and heteroduplex forms of mini-TAR are indicated by m and hd, respectively. (B) Percentage of heteroduplex in the presence of NCp9 or N1-419 at protein to nucleotide molar ratios of 1:28, 1:14 and 1:7. Black circles, NCp9; blue circles, 1-419. Experiments were repeated at least three times and the error bars indicate the standard error of the mean (SEM). ns, not statistically significant (unpaired two-samples t test).</figcaption></figure>

the first 20 s of the reaction and a slower one occurring during the 20-1800 s period of the reaction. Because the fast-kinetic step begins during the mixing time (10 s), the resolution of the first component could not be obtained at the highest ratios (1:15 and 1:8). The three parameters fast ($k_f$) and slow ($k_s$) annealing rates, and the fraction ($f$) of the fast component, were obtained by fitting the experimental data $P(t)$, the percentage of the DNA molecules annealed to RNA as a function of the time, to the general two-exponential expression (1) [61,62]:

$$P(t) = P_{\infty}(1 - f * \exp(-k_f t) - (1 - f) * \exp(-k_s t)) \quad (1)$$

$P_{\infty}$ is the ratio of DNA totally annealed to RNA and is assumed to be 1.0. Analysis of the annealing data with this bi-exponential equation has been made previously by several authors [60-62]. Our experiments with mutations in the stem or the apical loop

of mini-cTAR (Figure 1A) showed that the two types of mutations affected the kinetics (Supplementary Figure S2). The modified hairpins conserved a bi-exponential behavior, but the kinetics were largely affected with a diminution of 25-30% in the fluorescence intensity and the $k_{slow}$ rate. In addition, the decrease in hybrid formation was higher when the mutations were performed in the stem part (Supplementary Figure S2). These results suggest that the formation of the extended complex (the final hybrid duplex) can proceed by two pathways, one involving the apical loop (kissing complex) and another involving the stem ends (zipper complex). However, the pathway via the stem ends (the zipper model) was the leading pathway in our conditions, similar to the results obtained with the HIV-1 nucleocapsid protein [61]. Therefore, we used the model proposed and validated by Vo et al. [61,62] that is described in Materials and Methods. In this model,
<figure><img src="image_7.png" /><figcaption>**Figure 3.** Analysis of mini-TAR RNA-DNA annealing in the presence of N1-419, N31-210 or N240-419. The annealing assays were performed as described in Materials and Methods. (A) The samples were analyzed by electrophoresis on 15% polyacrylamide gels at 20 °C in the TBE buffer. The 5'-end labeled mini-TAR RNA alone was incubated in the absence of protein (lane C1) or in the presence of N1-419 (lane C3), N31-210 (lane C4) or N240-419 (lane C5). The 5'-end labeled mini-TAR RNA was incubated with mini-cTAR DNA in the absence of protein (lane C2) or in the presence of N1-419 (lanes 4–6), N31-210 (lanes 8–10) or N240-419 (lanes 12–14). The protein to nucleotide molar ratios were 1:28 (lanes 4, 8 and 12), 1:14 (lanes 5, 9 and 13) and 1:7 (lanes 3, 6, 7, 10, 11 and 14). The monomeric and heteroduplex forms of mini-TAR are indicated by m and hd, respectively. (B) Percentage of heteroduplex in the presence of N1-419, N31-210 or N240-419 at protein to nucleotide molar ratios of 1:28, 1:14 and 1:7. Blue circles, N1-419; orange squares, N31-210; purple triangles, N240-419. Experiments were repeated at least three times and the error bars indicate the standard error of the mean (SEM). Means determined with the truncated proteins were compared with those of the full-length. \*p < 0.05; \*\*p < 0.01; \*\*\*p < 0.005; \*\*\*\*p < 0.001; ns, not statistically significant (unpaired two-samples t test). (C) Percentage of heteroduplex in the absence or in the presence of protein at three different ratios. Orange lines, N31-210; purple lines, N240-419; black lines, no protein.</figcaption></figure>

the first step is the formation of an intermediary complex, in which the two hairpins are only partially annealed, which is involved in reversible equilibria with the free hairpins. The second step is the slow conversion of this intermediary complex to the fully extended duplex. The determination of the parameters associated with the first step of the reaction was strongly hampered by the duration of the mixing time (10 s) which prevented the observation of the beginning of the reaction. In contrast, the slow component was relatively easy to evaluate and was more useful because it really measures the ability of the nucleic acid chaperon to induce the formation of the extended duplex by destabilizing the secondary structures.

Figure 6 and Table 1 show the *f* (fraction) and *k*<sub>s</sub> (constant rate for the slow component) obtained from fitting kinetic data at three protein:nucleotide molar ratios (1:8, 1:15 and 1:29) and at least four

RNA concentrations. For the fast-kinetic phase, we extracted the fraction of intermediary complex (*f*) because it was not possible to extract accurate values of *k*<sub>fast</sub> for the reasons mentioned above. Interestingly, the data in Figure 6A show the dependence of *f* with RNA concentration and the protein:nucleotide molar ratio evolving from 1:29 to 1:8. For the second phase of the reaction (Figure 6B), the *k*<sub>slow</sub> constant appeared to be linear with the RNA concentration and a clear increase of the rate was observed when the protein:nucleotide molar ratio increased. Therefore, our results show that the SARS-CoV-2 nucleocapsid protein can increase the two phases of the annealing reaction. In particular, the *k*<sub>slow</sub> for the same RNA concentration was increased by a factor 2 from 1:29 to 1:15 and a factor 10 from 1:29 to 1:8 (considering the RNA concentration of 300 nM in Table 1).
<figure><img src="image_8.png" /></figure>

**Figure 4.** Nucleic acid annealing activity of N1-419 and its fragments. FRET efficiency ($E_{FRET}$) was measured as a function of protein concentration. The experiments were performed at pH 6.0 and 35 °C under the buffer conditions described in Materials and Methods. The mini-cTAR DNA was the labeled molecule bearing Fluorescein (F) and Tetramethylrhodamine (TMR) at its extremities, as indicated in the scheme and was at a concentration of 50 nM, while mini-TAR RNA was at a concentration of 300 nM. The data for the different proteins are shown by circles of various colors. The amino acid numbering has been indicated to identify the fragments. Data represent the mean ± SD of at least three independent experiments.

<figure><img src="image_9.png" /></figure>

**Figure 5.** Kinetics of mini-TAR DNA-RNA annealing in the presence of N1-419. The experiments were performed at a temperature of 35 °C and at pH 6.0 using the phosphate buffer solution described in Materials and Methods. The labeled mini-cTAR DNA was at a concentration of 50 nM and different concentrations of mini-TAR RNA were used. All the experiments were performed at a fixed protein to nucleotide ratio of 1:29. The signal corresponds to the spectra at 519 nm obtained in the presence of the protein from which the spectra of the two nucleic acids in the absence of the protein have been subtracted and converted in fluorescence arbitrary units.

It is interesting to compare these results with experiments performed with similar oligonucleotides and the HIV-1 nucleocapsid protein (NCp7), considered as one of the best nucleic acid chaperon. Note that the experiments with NCp7 were performed at 37 °C and pH 7.5 because at pH 6 its nucleic acid chaperone

activity is not optimal [70]. To compare the rate constants between different studies, we calculated the microscopic constant $k_2$ for the second phase of the kinetic that is not related to the concentrations used (see Materials and Methods). In a previous study, the $k_2$ value for NCp7 has been estimated to be $8 * 10^{-3}$ s$^{-1}$ at the 1:15 ratio [61] in experi-
<figure><img src="image_10.png" /><figcaption>**Figure 6.** RNA concentration and protein:nucleotide ratio dependence of mini-TAR DNA-RNA annealing parameters in the presence of N1-419. The parameters (fraction (f) and $k_{slow}$) have been obtained by fitting the FRET data as described in Materials and Methods. The labeled mini-cTAR DNA was used at a fixed concentration of 50 nM and the mini-TAR RNA concentration varied. The protein to nucleotide ratios were maintained fixed at values of 1:29 (orange), 1:15 (red) and 1:8 (green). (A) The fraction of intermediary complexes is given according to the RNA concentration and at the three fixed protein:nucleotide molar ratios; (B) The rate constant of the slow process ($k_{slow}$) is given according to the RNA concentration and at the three fixed protein:nucleotide molar ratios. Data represent the mean ± SD of at least three independent experiments.</figcaption></figure>

<table><caption>Table 1 Parameters of mini-TAR DNA-RNA annealing in presence of N1-419 deduced from the fit of the experimental data obtained at 35 °C, pH 6.0 using the buffer conditions described in Materials and Methods. The experimental conditions are the same as those described in Figure 3. The fraction (f value) and $k_{slow}$ constant are given at each protein to nucleotide molar ratio and RNA concentration.</caption><thead><tr><th>Protein to nucleotide ratio</th><th>RNA concentration</th><th>f value</th><th>k<sub>slow</sub> (s<sup>-1</sup>)</th></tr></thead><tbody><tr><td rowspan="5">Ratio 1:29</td><td>RNA 50 nM</td><td>0.17 ± 2.7 * 10<sup>-2</sup></td><td>1.1 * 10<sup>-4</sup> ± 7.0 * 10<sup>-5</sup></td></tr><tr><td>RNA 100 nM</td><td>0.18 ± 1.7 * 10<sup>-2</sup></td><td>1.3 * 10<sup>-3</sup> ± 1.3 * 10<sup>-4</sup></td></tr><tr><td>RNA 150 nM</td><td>0.21 ± 3.5 * 10<sup>-2</sup></td><td>1.5 * 10<sup>-3</sup> ± 1.8 * 10<sup>-4</sup></td></tr><tr><td>RNA 300 nM</td><td>0.26 ± 5.5 * 10<sup>-2</sup></td><td>2.9 * 10<sup>-3</sup> ± 2.0 * 10<sup>-4</sup></td></tr><tr><td>RNA 400 nM</td><td>0.39 ± 1.0 * 10<sup>-2</sup></td><td>4.0 * 10<sup>-3</sup> ± 4.0 * 10<sup>-5</sup></td></tr><tr><td rowspan="4">Ratio 1:15</td><td>RNA 50 nM</td><td>0.19 ± 4.5 * 10<sup>-4</sup></td><td>1.5 * 10<sup>-3</sup> ± 4.7 * 10<sup>-5</sup></td></tr><tr><td>RNA 100 nM</td><td>0.19 ± 4.6 * 10<sup>-3</sup></td><td>2.7 * 10<sup>-3</sup> ± 3.0 * 10<sup>-4</sup></td></tr><tr><td>RNA 150 nM</td><td>0.26 ± 3.6 * 10<sup>-2</sup></td><td>5.0 * 10<sup>-3</sup> ± 8.8 * 10<sup>-4</sup></td></tr><tr><td>RNA 300 nM</td><td>0.51 ± 7.4 * 10<sup>-2</sup></td><td>8.2 * 10<sup>-3</sup> ± 5.6 * 10<sup>-3</sup></td></tr><tr><td rowspan="4">Ratio 1:8</td><td>RNA 50 nM</td><td>0.47 ± 1.3 * 10<sup>-2</sup></td><td>2.8 * 10<sup>-3</sup> ± 2.1 * 10<sup>-4</sup></td></tr><tr><td>RNA 100 nM</td><td>0.43 ± 2.0 * 10<sup>-2</sup></td><td>7.4 * 10<sup>-3</sup> ± 3.0 * 10<sup>-4</sup></td></tr><tr><td>RNA 150 nM</td><td>0.51 ± 1.1 * 10<sup>-2</sup></td><td>1.3 * 10<sup>-2</sup> ± 3.9 * 10<sup>-4</sup></td></tr><tr><td>RNA 300 nM</td><td>0.62 ± 1.2 * 10<sup>-1</sup></td><td>4.0 * 10<sup>-2</sup> ± 1.0 * 10<sup>-2</sup></td></tr></tbody></table>

ments with similar oligonucleotides. In the laboratory we obtained $4 * 10^{-3}$ s$^{-1}$ with the same oligonucleotides as those used in the present study (results not shown), while the values obtained for SARS-CoV-2 for mini-TAR and mini-cTAR at 1:15 is $1.6 * 10^{-2}$ s$^{-1}$. The N protein seems therefore more efficient than NCp7 under similar conditions.

We determined the kinetics of mini-TAR DNA-RNA annealing in the presence of the N protein (N1-419) and three derived forms (double mutant

R203K/G204R, fragments N31-210 and N240-419). As shown by the fits in Figure 7 and the Table S1, the fragments N31-210 and N240-419 have approximately a $k_{slow}$ 3-fold and 7-fold lower than N1-419, respectively. Therefore, the activity of N31-210 appeared better than that of N240-419. These data agree with the results presented in Figure 4 showing that the N31-210 and N240-419 fragments are less efficient than N1-419 relative to mini-TAR DNA-RNA annealing and that
<figure><img src="image_11.png" /></figure>

**Figure 7.** Kinetics of mini-TAR DNA-RNA annealing in the presence of N1-419 (wt or mutant R203K/G204R forms) or its fragments (N31-210 and N240-419). The experiments were performed at a temperature of 35 °C and at pH 6.0 using the phosphate buffer solution described in Materials and Methods. The labeled mini-cTAR DNA concentration was 50 nM and the mini-TAR RNA concentration was 150 nM. The experiments were performed at a protein to nucleotide molar ratio of 1:15. The signal corresponds to the spectra at 519 nm obtained in the presence of the protein from which the spectra of the two nucleic acids in the absence of the protein have been subtracted and converted in fluorescence arbitrary units.

there is a significant difference between N31-210 and N240-419. They also demonstrate the relevance of the kinetic constants under the conditions chosen to describe the efficiency of the annealing process. The mutant protein showed a similar efficiency relative to the wild-type protein (Figure 7).

In addition, we determined by FRET the effect of pH on the nucleic acid chaperone activity of N1-419. More precisely, the experiments were performed with the same buffers and temperature at three different pH (Figure 8). Interestingly, the nucleic acid chaperone activity strongly increased when the pH decreased and was optimal at low pH (6.0).

To show that N1-419 can promote the annealing of longer stem-loops, we performed another set of experiments using the cTAR and TAR hairpins that are 59 nucleotides long (Figure 1) [62]. The kinetic data obtained with these stem-loops were deduced from Figure 9. The $k_{slow}$ and $k_2$ at a protein to nucleotide molar ratio of 1:29 were $2.7 * 10^{-3} s^{-1}$ and $4.1 * 10^{-3} s^{-1}$, respectively (Table S2). Under the same conditions, the values for annealing of mini-cTAR to mini-TAR were $2.9 * 10^{-3} s^{-1}$ and $1.1 * 10^{-2} s^{-1}$, indicating that annealing was only slightly faster with small stem-loops. The $k_2$ value was close to $6 * 10^{-3} s^{-1}$ when the annealing reaction was performed with the small stem-loops and NCp7 at a protein to nucleotide molar ratio of 1:15 (not shown). Taken together, our results demonstrate the high annealing potential of the SARS-CoV-2N protein. Interestingly, the mutated N protein bearing two mutations appeared more efficient than the wild-type protein in annealing the long substrates ($k_{slow}$ is $4.1 * 10^{-3} s^{-1}$).

Finally, to investigate in detail the nucleic acid chaperone activity of N1-419, we determine its ability to aggregate nucleic acids. We performed the experiments under the same conditions as the EMSA experiments. As described in the Materials and Methods section, we added a step of centrifugation to separate the aggregates from the soluble nucleic acids. We also used NCp9 as a positive control because its aggregation properties are well-known [71,72]. Figures 10 and S3 display the results of aggregation assays. The percentage of mini-TAR RNA aggregated is proportional to the protein concentration (Figure 10A). These data are consistent with the notion that nucleic acid aggregation is an important component of protein N's chaperone function. Interestingly, we observed that its aggregation activity at the 1:14 and 1:7 ratios is significantly higher than that of NCp9 (Figure 10B).

## Destabilizing effect of N protein and fragments on small double-stranded RNA duplexes

We analyzed the ability of the N protein and of three of its fragments to destabilize small RNA duplexes corresponding to Transcription Regulating Sequences (TRS) of SARS-CoV-2. These sequences are involved in the discontinuous transcription occurring during the synthesis of negative-sens subgenomic RNAs (sgRNAs). The chosen sequence corresponds to the TRS leader sequence located at the 5'-end of the viral genome [73,74]. The experiments were performed at 10 °C with the five proteins (Figure 1B), and an RNA duplex of 9 nucleotides possessing a
<figure><img src="image_12.png" /></figure>

**Figure 8.** Nucleic acid annealing activity of N1-419 at different pH. FRET efficiency ($E_{FRET}$) was measured as a function of protein concentration. The experiments have been performed at 35 °C under the buffer conditions described in Materials and Methods. The labeled mini-cTAR DNA concentration was 50 nM and the mini-TAR RNA concentration was 300 nM. The data for the full-length protein at different pH are shown by circles of various colors. Data represent the mean ± SD of at least three independent experiments.

<figure><img src="image_13.png" /></figure>

**Figure 9.** Kinetics of TAR DNA-RNA annealing in the presence of N1-419 and N1-419mut. The experiments were performed at a temperature of 35 °C and at pH 6.0 using the phosphate buffer solution described in Materials and Methods. The labeled cTAR DNA concentration was 50 nM and the TAR RNA concentration was 300 nM. The experiments were performed at a protein to nucleotide molar ratio of 1:29. The signal corresponds to the spectra at 519 nm obtained in the presence of protein from which the spectra of the two nucleic acids in the absence of the protein have been subtracted and converted in fluorescence arbitrary units. In blue are shown the data of TAR DNA-RNA annealing without protein.

Tm of around 20 °C. The two oligonucleotides forming the RNA duplex bear the Cy3 and Cy5 fluorescent probes as described in Materials and Methods. We added each protein at increasing concentrations (Supplementary Figure S4). The patterns of the spectra for the three truncated proteins were different. In the presence of increasing concentrations of N31-210, the Cy3 signal at 560 nm increased, whereas the Cy5 signal at 665 nm strongly decreased. This latter behavior was expected for a FRET effect. However, for the

highest protein concentration, quenching effects of N31-210 upon Cy3 signal complicated the evaluation of FRET efficiency. In the presence of increasing concentrations of N1-260, the Cy3 signal increased while the Cy5 signal slightly decreased. In the presence of increasing concentrations of N240-419 as well as for N1-419, the signal variations are particularly weak for Cy3 and Cy5, showing the inability of N240-419 protein to melt the small RNA duplex. Similarly, N1-419 and its mutated version N1-419 mut (R203K/G204R) did
<figure><img src="image_14.png" /><figcaption>**Figure 10.** Analysis of mini-TAR RNA aggregation. The aggregation assays were performed as described in Materials and Methods. (A) The percentage of mini-TAR RNA aggregated was determined in the absence of protein or in the presence of NCp9 or N1-419 at protein to nucleotide molar ratios of 1:28, 1:14 and 1:7. Black circles, NCp9; blue circles, 1–419. Experiments were repeated at least three times and the error bars indicate the standard error of the mean (SEM). Means determined with the proteins were compared with that of the control (no protein). (B) Means determined with NCp9 were compared with those of N1-419. **p** < 0.01; ***p** < 0.005; ****p** < 0.001; ns, not statistically significant (unpaired two-samples *t* test).</figcaption></figure>

not produce significant variations of Cy3 and Cy5 fluorescence signals.

The quenching effect of proteins on the Cy3 fluorescence signal (performed in the absence of Cy5 fluorophore) suggest, in agreement with several previous reports, it exist a significative binding of the N protein on the RNA under the single-stranded or double-stranded forms [27]. To confirm these results, we analyzed by fluorescence anisotropy the binding of the five proteins to the TRS oligonucleotide under single-stranded or double-stranded forms. The two full-length proteins (wild-type and mutant), N31-210 and N1-260 bound the oligonucleotides with significant affinities (Figure S5 and Table S3), while this was not the case for N240-419. N31-210 and N1-260 displayed approximately 3-fold and 4-fold higher affinity for single-stranded relative to double-stranded oligonucleotides, respectively. These results agree with a previous study [27].

Finally, our experiments showed that the N31-210 fragment has a higher destabilizing activity than the four other proteins. In addition, our binding study suggests that the single-stranded parts of the RNA duplex resulting from the destabilizing activity of the N protein are bound subsequently by the protein with a significant affinity.

## Discussion

Despite numerous recent works on the ability of the SARS-CoV-2 N protein to form liquid-liquid phase separation (LLPS) in the presence of RNA [13,20,41,42,53,75], the studies on the ability of this

protein to act on RNA-RNA annealing or nucleic acid melting have been limited [24,76,77].

Here, we analyzed the aggregative, strand-annealing, and RNA-melting properties of the SARS-CoV-2 N protein and several of its fragments. We used a model system involving DNA and RNA hairpins derived from the HIV-1 *trans*-activation response (TAR) element to analyze the aggregative and annealing activities. Its use permits us to show that the annealing and aggregative activities of the SARS-CoV-2N protein are similar or higher to those of the HIV-1 nucleocapsid proteins NCp7 and NCp9 (Figures 2 and 10) that are both excellent nucleic acid chaperons [66,67,78–80].

We used two different methodologies to measure the annealing properties of the N protein: (i) annealing experiments with oligonucleotides in which the DNA molecule bore chromophores at each extremity and the annealing was revealed with the loss of the FRET transfer and (ii) Electrophoretic Mobility Shift Assays (EMSA) in which RNA carries a fluorescent dye; the heteroduplex product and the RNA monomer were identified through different mobilities in the polyacrylamide gel. The results obtained with the two methodologies led to the same conclusion: the N protein possesses a strong nucleic acid chaperone activity, as efficient as NCp7 and NCp9.

FRET experiments with fragments of the N protein showed that the most efficient fragments are those containing the NTE, RBD and LINK domains (Figure 1B, N1-260 and N31-210), while N240-419 is poorly efficient (Figure 4). However,
the full-length protein appeared to possess a better annealing activity than any of the fragments. A similar study [16] performed with the N protein of transmissible gastroenteritis virus (TGEV) and its fragments showed that the most efficient fragments for the annealing activity were those containing the central linker (LINK) and the N-terminal (NTD) parts. In addition, these fragments were found more efficient than the full-length protein for the annealing activity [16]. The difference in properties between the SARS-Cov-2 and TGEV nucleocapsid proteins is puzzling. A significant difference is that in our experiments, the two complementary nucleic acids possessed stable secondary structures, while this was the case for only one of the complementary nucleic acids in the experiments with the TGEV N protein [16]. Another hypothesis is that the fragments of the SARS-CoV-2N protein, in contrast to those of the TGEV N protein, partially reflect the functional domains. However, our results are qualitatively similar to those obtained with the TGEV N protein since the fragments containing the central linker and the N-terminal domains are the most efficient, and the one containing the C-terminal domain displays poor annealing activities [15,16].

The destabilizing experiments were performed with small RNA duplexes of 9 or 11 base pairs. The FRET effect associated with the duplex destabilization was not easy to measure due to the significative binding of the protein close to the fluorescent probes, leading either to exaltation of fluorescein or quenching of the Cy3 probe. In addition, we measured the affinity binding of proteins to single-stranded and double-stranded RNAs using anisotropy fluorescence spectroscopy. The N31-210 and N1-260 fragments, containing the RBD and SR (part 176–206 of the LINK domain [81]) domains (Figure 1B), were the most efficient in destabilizing the mini RNA duplex. In contrast, the full-length proteins (wild-type and mutant) and the N240-419 fragment were poorly efficient. These results are consistent with previous studies showing that the NTD, SR, and linker domains of the N proteins of coronaviruses are the most efficient in destabilizing RNA duplexes [24,57,58]. Although N1-260 contains the totality of the NTE, RBD, SR, and LINK domains, this fragment is less efficient than N31-210, containing the RBD and SR domains but truncated forms of the NTE and LINK domains. It is possible that the NTE domain interacts with the RBD domain and modifies the RNA binding properties of RBD and SR domains. Consistent with this hypothesis, a previous study showed that in the context of an NTE-RBD construct, significant interactions were observed between the NTE and RBD domains [20]. An important result is that the full-length protein is inefficient in destabilizing the small TRS-cTRS RNA duplexes, possibly because these oligonucleotides are too short. Interestingly, a recent study also showed the weak ability of the full-length protein to destabilize RNA duplexes relative to active helicases such as nsp13, the SARS-CoV-2 helicase [76].

Two recent studies compared the nucleic acid binding properties of SARS-CoV-2N protein to its fragments. Using fluorescence anisotropy, it was found that the N protein possesses a higher affinity for single-stranded RNA than for double-stranded RNA with a difference of a factor 7–10 [27]. Our annealing experiments showed that N240-419, containing the dimerization and CTE domains, is inefficient in annealing complementary stem-loops (Figure 4). In contrast, Morse et al. [77] showed that the protein fragment containing these domains possesses a significant compaction activity but at a slow rate relative to the full-length protein. Using two annealing assays, we showed that the full-length protein is required for efficient annealing of complementary hairpins (Figures 3 and 4). The absence of the dimerization domain in N31-210 and N1-260 but its presence in N1-419 suggests that dimerization of the full-length protein plays an essential role in the annealing reaction. The fact that N31-210 is more efficient than N240-419 in the FRET assays but not in the EMSA assays could be due to the different experimental conditions used. Indeed, the pH and buffer can change the oligomerization and LLPS properties of the protein [82].

An interesting question is to know what oligomeric forms of the N protein are involved in the annealing process. Oligomerization depends on the presence of RNA and probably also on its size and nature (single-stranded vs double-stranded, presence of secondary structures, base composition) [52,53,83–85]. Carlson and Morgan proposed [53] a model in which the dimer of the N protein bound to one RNA stem-loop (80 nt) assembles iteratively to constitute a large viral RNP ensemble containing a fixed ratio between dimers and the number of stem-loops. Consistent with this model, our results showed that the full-length N protein annealed efficiently small stem-loops (29 or 59 nt), but it was not the case for the N1-260 and N31-210 fragments lacking the dimerization domain.

Interestingly, we established that pH is an important parameter for the annealing efficiency (Figure 8). Previous studies showed that protein-protein interactions of SARS-CoV-2N protein are enhanced at low pH, suggesting that the annealing reaction is more efficient at pH 6 because this condition favors protein oligomerization [41,47,82].

A striking result from our study is the very efficient ability of the full-length N protein to destabilize and anneal stable RNA stem-loops as mini-TAR or TAR. Our experiments suggest that the SARS-CoV-2N protein is as efficient as HIV-1 nucleocapsid proteins (NCp7 and NCp9) that are powerful nucleic acid chaperons. One can wonder at what level of the viral replication cycle, such
activity should be involved. The first hypothesis is the involvement of the N protein in the continuous and discontinuous transcription [16,58]. Interestingly, the TRS-body elements located near the ORF of the structural proteins are nearly always inside a stable hairpin (7 on a total of 9 TRS-B) [74,86]. Another hypothesis is that the nucleic acid chaperone activity of the N protein is required for the destabilization of numerous secondary structures of the SARS-Cov-2 genome [86,87], i.e. just before the entry of the genome inside the pores of the double membrane vesicle [88,89] or at the step of genome packaging leading to the formation of the nucleocapsid complex between viral RNA and the N protein [90]. At these steps, the disentanglement of the RNA genome by the N protein and associated proteins should be essential to increase the speed of the replication cycle. Relative to this last point, several recent publications underline the involvement of the N protein in an increase in the replication rate [3,90–92].

In conclusion, we showed that the SARS-CoV-2N protein exhibits significant nucleic acid chaperone activity, similar to that of HIV-1 nucleocapsid proteins. This activity is more efficient with the full-length than with truncated N proteins. Our results support the notion that the RBD domain plays a key role in nucleic acid chaperone activity, and the C-terminal part of the N protein increases this activity by allowing protein dimerization. Moreover, the ability of the N protein to anneal complementary hairpins increases at acidic pH, which favors protein oligomerization. The N protein's chaperone activity is likely involved in the transcription and packaging steps of SARS-Cov-2.

# Materials and Methods

## Proteins

NCp7(1–72) (referred to as NCp9) was synthesized and purified as previously described [93]. NCp7(1–72) possesses an additional amino acid at its c-terminus in comparison to NCp7(1–71) which is the natural NCp9 [67,93,94].

The gene encoding SARS-CoV-2 nucleocapsid protein (Gene ID: 43740578) was cloned into the protein expression vector pET15b-SUMO (Takara), which was used to transform the C2566H *E. coli* strain (New England Biolabs). When the culture reached the early stationary phase (OD~600~ = 0.5–0.6) at 37 °C, 0.3 mM IPTG (Isopropyl-β-D-thiogalactopyranoside) was added, and the protein expression was induced at 18 °C over 15 h. After harvesting the cells, pellets were resuspended in lysis buffer (20 mM Tris–HCl pH 7.5, 300 mM NaCl, 10 mM Imidazole, and 5% glycerol (v/v)). Cells were broken with an ultra-high-pressure cell disrupter (JNBIO) and then further sonicated 3 times to shear DNA. After centrifugation at 12,000 rpm for 40 min, the supernatants were filtered through a 0.45-µm filter

and loaded onto the Ni-NTA column (QIAGEN). After washing with wash buffer (20 mM Tris–HCl, pH 7.5, 300 mM NaCl, 20 mM Imidazole, and 5% glycerol (v/v)), the SUMO fusion protein was eluted with elution buffer (20 mM Tris–HCl, pH 7.5, 300 mM NaCl, 500 mM Imidazole and 5% glycerol (v/v)). The SUMO tag was cleaved with SUMO protease at 4 °C overnight, and the protein was further purified by SP Sepharose Fast Flow (GE Healthcare) and Q Sepharose Fast Flow (GE Healthcare), respectively. The protein was finally purified by gel-filtration chromatography on HiLoad Superdex 200 pre-equilibrated with 20 mM Tris–HCl, pH 7.5, and 300 mM NaCl. After the final step, the purified proteins were about 95% pure as determined by SDS-PAGE and stored at –80 °C. All mutants and truncated proteins were expressed and purified following a similar procedure described above.

## Oligonucleotides

RNA and DNA oligonucleotides corresponding to the HIV-1 TAR, mini-TAR, cTAR and mini-cTAR stem-loops (Figure 1A), and complementary RNA oligonucleotides forming the TRS RNA duplex were purchased from Eurogentec (Belgium). In the EMSA and aggregation experiments, we used the mini-cTAR DNA and the mini-TAR RNA labeled at its 5'-terminus by Cy®5.5 that is a fluorescent dye (absorbance, 675 nm; emission 694 nm). In the annealing assays analyzed by fluorescence spectroscopy, two fluorescent probes were grafted at the two extremities of the cTAR/mini-cTAR DNA: 5/6-carboxytetramethylrhodamine (TMR) at the 5'-end and 5/6-carboxyfluorescein (F) at the 3'-end. In the destabilization assays, we used two complementary RNA oligonucleotides: ssTRS(+) (5'-CUAAACGAA-3') labeled at its 5'-end by the Cy3 fluorophore and ssTRS(−) (5'-UUCGUUUAG-3') labeled at its 3'-end by the Cy5 fluorophore.

## Fluorescence spectroscopy

All the fluorescence experiments were performed in 50 mM potassium phosphate buffer with 100 mM KCl, pH 6.0. In particular experiments, the pH was modified to evaluate its impact on the annealing reaction. For the kinetic measurements, the DNA oligonucleotide concentration was 50 nM, the RNA concentration generally varied from 50 to 400 nM and the protein was added at a protein to nucleotide molar ratio that varied from 1:6 to 1:29.

Förster Resonance Energy transfer (FRET) experiments were performed on a Fluoromax spectrofluorometer equipped with a quartz cell of 1.0 cm optical path length and equipped with a temperature control system. A bandwidth of 4 nm was set both for excitation and emission. FRET measurements were carried out with DNA
oligonucleotides labeled at the 5' and 3' ends with the acceptor probe TMR and the donor probe Fluorescein, respectively. The donor was excited at 480 nm and the emission spectrum was collected in the 500-700 nm range, observing the FRET efficiency (E) of the acceptor probe at the maximum emission at 519 nm. For the experiments with Cy3-Cy5 pair that possess the following properties: ($\lambda_{ex}$ = 520 nm for Cy3 excitation, $\lambda_{em}$ = 550-600 nm for Cy3 emission, and with $\lambda_{ex}$ = 635 nm for Cy5 excitation, $\lambda_{em}$ = 65 5-695 nm for Cy5 emission).

For the measurement of the FRET efficiency, the DNA oligonucleotide concentration was set to 50 nM while the final concentration of RNA was set to 300 nM and the final protein concentration was in the range 10-600 nM. The FRET efficiency was measured after initial mixing of DNA with RNA and protein and 15 min of delay to ensure the stabilization of the signal. The FRET efficiency was calculated from $E = 1 - \frac{F_{DA}}{F_D}$, where $F_D$ and $F_{DA}$ are the fluorescence intensities of the donor Fluorescein in the absence and in the presence of the acceptor (TMR), respectively, E being the FRET efficiency for the oligonucleotide probes with the pair Cy3-Cy5. Experiments with addition of protein upon the single and double stranded TRS9_Cy3 (in which the complementary strand did not possess the Cy5 partner) show that the protein alone cause a significative quenching of the Cy3 signal. These later experiments were therefore used to correct the intensity of Cy3 signal and determine the FRET efficiency.

Fluorescence anisotropy assays experiments were performed on the Fluoromax spectrofluorometer. Anisotropy measurements were carried out with the single-stranded TRS9ss (5'-U U C G U U U A G 3') and double-stranded TRS9ds (5'-U U C G U U U A G 3' plus 5'C U A A A C G A A 3') RNA oligonucleotides described. Excitation and emission wavelengths were set to 480 nm and 516 nm for TRS9ss and to 480 nm and 512 nm for TRS9ds. The RNA oligonucleotide at a concentration of 50 nM was incubated in the same buffer than for the FRET experiments, and the protein was added increasingly, at each protein concentration fluorescence anisotropy was measured.

## EMSA experiments

The annealing assay (final concentrations: 36 mM NaCl, 0.12 mM MgCl₂, 0.012 mM ZnCl₂, 5 mM DTT and 60 mM Tris-HCl, pH 7.5) was carried out in a final volume of 10 µL. The experimental steps are described below. Two pmol of the 5'-end labeled mini-TAR RNA alone or with 2 pmol of mini-cTAR DNA in 6.75 µL of water were heated at 90 °C for 2 min and chilled for 2 min on ice. Then, 1.25 µL of renaturation buffer (240 mM NaCl, 0.8 mM MgCl₂, 0.08 mM ZnCl₂, 40 mM DTT and 400 mM Tris-HCl, pH 7.5) was added and the sample was

incubated at 25 °C for 10 min. Two µL of NC buffer (30 mM NaCl, 0.1 mM MgCl₂, 0.01 mM ZnCl₂ and 50 mM Tris-HCl, pH 7.5) with or without protein were added. The protein to nucleotide molar ratios were 1:28, 1:14 or 1:7. These ratios refer to total nucleotide concentration. The reaction mixture was then incubated at 37 °C for 10 min. At the end of incubation, 2 µL of stop solution (4.5% SDS, 25 mM EDTA and 1.5 µg/µL of proteinase K) was added before an incubation of 15 min at 25 °C. Then, the assay was phenol/chloroform extracted and the aqueous phase was mixed with 4 µL of loading buffer (50% w/v glycerol, 0.05% w/v bromophenol blue, 0.05% w/v xylene cyanol). The samples were analyzed by 15% polyacrylamide gel (37.5:1 (w/v), acrylamide/bisacrylamide) at 20 °C in the TBE buffer (89 mM Tris-borate (pH 8.3), 2 mM EDTA). After the electrophoresis, the fluorescence of the monomeric (m) and heteroduplex (hd) forms of mini-TAR RNA were visualized and quantified using the Amersham™ Typhoon™ 5 device and Image-Quant software (Cytiva, France). The percent of heteroduplex was determined as $100 \times (hd/(hd + m))$.

## Aggregation assay

The aggregation assay (final concentrations: 36 mM NaCl, 0.12 mM MgCl₂, 0.012 mM ZnCl₂, 5 mM DTT and 60 mM Tris-HCl, pH 7.5) was carried out in a final volume of 48 µL. The experimental steps are described below. The Cy® 5.5 mini-TAR RNA (9.6 pmol) with 9.6 pmol of mini-cTAR DNA in 32.4 µL of water were heated at 90 °C for 2 min and chilled for 2 min on ice. Six µL of renaturation buffer (240 mM NaCl, 0.8 mM MgCl₂, 0.08 mM ZnCl₂, 40 mM DTT and 400 mM Tris-HCl, pH 7.5) was added and the sample was incubated at 25 °C for 10 min. Then, 9.6 µL of NC buffer (30 mM NaCl, 0.1 mM MgCl₂, 0.01 mM ZnCl₂ and 50 mM Tris-HCl, pH 7.5) with or without protein were added. The protein to nucleotide molar ratios were 1:28, 1:14 or 1:7. These ratios refer to total nucleotide concentration. The reaction mixture was then incubated at 37 °C for 10 min. At the end of the incubation time, the aggregation assays were centrifuged at 12,000 rpm, 4 °C for 20 min. This centrifugation step, allowing separation of soluble nucleic acids from aggregates [61,62,95,96], was adapted from Vo et al. 2006. The aggregates formed by the nucleic acids and the proteins sediment to the bottom of the tube upon centrifugation, whereas the soluble nucleic acids are distributed homogeneously throughout the upper part of the tube. The upper part of each assay (8 µl) was removed carefully and transferred to another tube before to be analyzed by fluorescence. The fluorescence spectra of Cy® 5.5 were recorded between 685 and 800 nm by exciting the sample at 670 nm. Fluorescence emission spectra resulted
from at least three accumulations, each corrected for buffer background contribution. The spectra were obtained at 25 °C. For each sample, the fluorescence intensity at 696 nm corresponding to the soluble nucleic acids was determined. The fluorescence intensity of control (mini-TAR + mini-cTAR in the absence of protein) was arbitrarily set to represent 100% of soluble nucleic acids. For each sample, the percentage of soluble nucleic acids was normalized to the control. The percentage of aggregated nucleic acids corresponded to 100 minus the percentage of soluble nucleic acids.

### Model used for the extraction of kinetic data

The model displaying two kinetic phases has been developed by Vo et al. [61,62], it describes the kinetics of mini-TAR RNA-DNA and TAR RNA-DNA annealing in the presence of the N protein:

$$ TAR + cTAR \underset{k_{-1}}{\rightleftharpoons} TARcTAR \underset{k_{-2}}{\rightleftharpoons} E.D $$

The first bimolecular step leads to the formation of the annealing intermediate (RNA/DNA)* with the forward association rate constant $k_1$ (R is the RNA concentration) and the intermediate dissociation rate $k_{-1}$. The annealing intermediate (RNA/DNA)* is converted into the RNA-DNA extended duplex, the fully annealed hybrid, in a monomolecular step characterized by the strand exchange rate $k_2$ and the reverse reaction $k_{-2}$. In our treatment, the reverse reaction was neglected. We used Prism (https://www.graphpad.com/scientific-software/prism/) and R program (https://www.r-project.org/) to extract the $k_{fast}$, $k_{slow}$ and f values from the experimental curves in various conditions. We noted that the $k_{fast}$ could not be extracted when the protein to nucleotide molar ratio is too high (1:8 and 1:15) in the presence of high RNA concentrations. Under these conditions the fast process occured largely during the deadtime of the experiment (10 s). The microscopic constant $k_2$ was determined using the relation $k_2 = k_{slow}/f$, where f is the fraction of the fast component [61,62].

To check if the experimental data were compatible with the hypothesis of the same value for $k_1$, $k_{-1}$ and $k_2$ at a constant protein to nucleotide molar ratio, we simulated the kinetics at a constant protein to nucleotide molar ratio with all the RNA concentrations. For this, we used the Scipy Python suite of programs and the odeint (integrator for ordinary differential equations) that allow us to use the experimental concentrations and do not rely on pseudo-first order approximation. In these simulations, we assigned an arbitrary FRET value to the annealing intermediate, the value of 0.5 gave the best fit, this seems reasonable as the intermediary species is mostly the "zipper complex". In this

latter form, the two fluorophores are expected to be closer relative to the extended duplex.

### CRediT authorship contribution statement

**Loussine Zargarian:** Data curation, Formal analysis, Methodology, Investigation, Writing – review & editing. **Xia Ai:** Investigation. **Ze-Yu Song:** Investigation. **Lara Perez-Gorgol:** Investigation. **Esteban Couprie-Diaz:** Investigation. **Philippe Fossé:** Methodology, Supervision, Writing – review & editing, Conceptualization, Writing – original draft, Formal analysis, Data curation. **Xu-Guang Xi:** Supervision, Validation, Writing – review & editing, Resources, Methodology. **Olivier Mauffret:** Funding acquisition, Project administration, Writing – original draft, Conceptualization, Supervision, Formal analysis, Writing – review & editing, Data curation.

### DECLARATION OF COMPETING INTEREST

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Appendix A. Supplementary material

Supplementary material to this article can be found online at https://doi.org/10.1016/j.jmb.2025.169312.

Received 27 February 2025;
Accepted 23 June 2025;
Available online 30 June 2025

**Keywords:**
SARS-CoV-2;
N protein;
nucleocapsid;
chaperon protein;
stem-loop structures

† Equal contribution.

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<table><caption>Supplementary Table S1.</caption><thead><tr><th>Protein to<br>nucleotide<br>ratio</th><th>RNA<br>concentration</th><th>Protein</th><th>f value</th><th>k<sub>slow</sub> (s<sup>-1</sup>)</th></tr></thead><tbody><tr><td rowspan="4">1:15</td><td rowspan="4">150nM</td><td>N1-419</td><td>0.26 ± 3.6*10<sup>-2</sup></td><td>5.0*10<sup>-3</sup> ± 8.8*10<sup>-4</sup></td></tr><tr><td>N1-419mut</td><td>0.30 ± 2.2*10<sup>-2</sup></td><td>2.6*10<sup>-3</sup> ± 1.8*10<sup>-4</sup></td></tr><tr><td>N31-210</td><td>0.22 ± 1.1*10<sup>-3</sup></td><td>1.5*10<sup>-3</sup> ± 3.8*10<sup>-4</sup></td></tr><tr><td>N240-419</td><td>0.30 ± 4.6*10<sup>-3</sup></td><td>6.9*10<sup>-4</sup> ± 1.1*10<sup>-4</sup></td></tr></tbody></table>
<table><thead><tr><th></th><th>Protein to<br>nucleotide<br>ratio</th><th>RNA<br>concentration</th><th>f value</th><th>k<sub>slow</sub> (s<sup>-1</sup>)</th><th>k<sub>2</sub> (s<sup>-1</sup>)</th></tr></thead><tbody><tr><td><strong>mini-cTAR/mini-<br>TAR</strong></td><td>1:29</td><td>300nM</td><td>0.26 ± 5.5*10<sup>-2</sup></td><td>2.9*10<sup>-3</sup> ± 2.0*10<sup>-4</sup></td><td>1.1*10<sup>-2</sup> ± 3.6*10<sup>-3</sup></td></tr><tr><td><strong>cTAR/TAR</strong></td><td></td><td></td><td>0.65 ± 5.3*10<sup>-2</sup></td><td>2.7*10<sup>-3</sup> ± 4.4*10<sup>-4</sup></td><td>4.1*10<sup>-3</sup> ± 2.0*10<sup>-4</sup></td></tr></tbody></table>

Supplementary Table S2.
<table><caption>Supplementary Table S3</caption><thead><tr><th>RNA concentration</th><th>Protein</th><th>Kd (M)<br>TRS9ds</th><th>Kd (M)<br>TRS9ss</th></tr></thead><tbody><tr><td rowspan="4">50nm</td><td>N1-419</td><td>2.1*10<sup>-7</sup> ± 4.2*10<sup>-8</sup></td><td>1.1*10<sup>-7</sup> ± 2.7*10<sup>-9</sup></td></tr><tr><td>N1-419mut</td><td>3.6*10<sup>-7</sup> ± 4.3*10<sup>-9</sup></td><td>2.2*10<sup>-7</sup> ± 1.4*10<sup>-8</sup></td></tr><tr><td>N1-260</td><td>1.6*10<sup>-7</sup> ± 2.8*10<sup>-8</sup></td><td>4.3*10<sup>-8</sup> ± 4.0*10<sup>-9</sup></td></tr><tr><td>N31-210</td><td>3.1*10<sup>-8</sup> ± 7.3*10<sup>-9</sup></td><td>9.3*10<sup>-9</sup> ± 3.7*10<sup>-9</sup></td></tr></tbody></table>
<figure><img src="image_15.png" /><figcaption>Supplementary Figure S1. Kinetics of mini-TAR DNA-RNA annealing in the presence of N1-419 (in blue) and absence of the protein (in red). The experiments were performed at a temperature of 35 °C and at pH 6.0 using the phosphate buffer solution described in Materials and Methods. The labeled mini-cTAR DNA concentration was 50 nM and the mini-TAR RNA concentration was 300 nM. The experiments with the protein were performed at a protein to nucleotide molar ratio of 1:29. The signal is the fluorescence intensity at 519 nm was converted in fluorescence arbitrary units.</figcaption></figure>

<figure><img src="image_16.png" /><figcaption>Supplementary Figure S2. Kinetics of mini-TAR DNA-RNA and of mini-TAR DNA mutants-RNA in the presence of N1-419. The experiments were performed at a temperature of 35 °C and at pH 6.0 using the phosphate buffer solution described in Materials and Methods. The labeled mini-cTAR DNA (wt or mutant) concentration was 50 nM and the mini-TAR RNA concentrations was 150 nM. The experiments were performed at a protein to nucleotide molar ratio of 1:15. Mini-cTAR is the wt form, in mini-cTAR-loop, two residues of the apical loop have been mutated; in mini-cTAR-stem, two residues of the stem have been modified. The signal corresponds to the spectra at 519 nm obtained in the presence of the protein from which the spectra of the two nucleic acids in the absence of the protein have been subtracted and converted in fluorescence arbitrary units.</figcaption></figure>
<figure><img src="image_17.png" /><figcaption>Supplementary Figure S3. Fluorescence analysis of mini-TAR RNA aggregation. The aggregation assays were performed as described in Materials and Methods. Fluorescence spectra of the Cy®5.5 mini-TAR RNA were determined in the absence of protein or in the presence of NCp9 or N1-419 at protein to nucleotide molar ratios of 1:28, 1:14 and 1:7. Black lines, NCp9; blue lines, N1-419; dashed lines, no protein.</figcaption></figure>

<figure><img src="image_18.png" /></figure>

<figure><img src="image_19.png" /><figcaption>Supplementary Figure S4. Fluorescence spectra of the 9-Cy3-TRSs(+)/9-Cy5-TRSs(-) RNA duplex (50 nM) in the absence and presence of N1-419, N1-419mut, N1-260, N31-210 or N240-419. The spectra were obtained at 10 °C with an excitation wavelength λ<sub>exc</sub> of 520 nm, that is the maximum absorption for Cy3, the emission is maximal for λ<sub>em</sub> = 562 nm for Cy3 and λ<sub>em</sub> = 670 nm for Cy5. The FRET efficiency is determined in function of the protein concentration. The buffer conditions are 50 mM potassium phosphate, 100 mM KCl, pH 6.0.</figcaption></figure>
<figure><img src="image_20.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_21.png" /></figure>

Supplementary Figure S5. Fluorescence polarization binding curves of N constructs to a TRS-9 nt RNA molecule bearing a fluorescein probe at the 3' end, either under double-stranded form (A) or single-stranded forms (B). The anisotropy values are indicated versus the concentrations of proteins expressed in M. The experiments were performed at a temperature of 10 °C and at pH 6.0 The buffer conditions are 50 mM potassium phosphate, 100 mM KCl, pH 6.0. The labeled TRS-9nt RNA concentration and RNA duplex was 50 nM. The color are related to the five used constructs.