Recently we have reported the discovery of novel miRNAspecific metal-independent aRNases, here called miRNases, capable of selective cleavage of biologically significant miRNAs (Patutina et al., 2017a), where we presented a design concept, synthesis, and full characterization of a panel of the bioconjugates at their discovery stage with the significant structural and functional variations. Each of those miRNases comprised an oligodeoxyribonucleotide complementary to the miRNA of interest as a recognition motif conjugated to peptide acetyl-[(LeuArg)2Gly]2 capable of a site-specific scission of phosphodiester bonds within the target miRNA molecule. That pilot study allowed us to compare *back-to-back* the ability of such conjugates to cleave the target miR-21 in a sequencespecific manner to distinguish successful structural candidates from inactive counterparts. One of the most efficient miRNases from that series (i.e. 5′-h-9/14) was used for preliminary evaluation of a biological activity in order to estimate the future antitumor potential of these catalytic bioconjugates. As the selected miRNase targeted to the oncogenic miR-21 (miR-21-miRNase) showed efficient sequence-selective cleavage, leading to inhibition of miR-21 in lymphosarcoma cells and suppression of tumor cell proliferation (Patutina et al., 2017a), we undertake here a comprehensive study of its serum stability and biological performance. We demonstrate that specific deactivation of miR-21 by miR-21-miRNase induces apoptosis in tumor cells, suppresses their invasive properties, and decreases the proliferative rate of tumor cells *in vitro*. We show here for the first time that a single treatment of tumor cells with miR-21 miRNase persisted to retard subsequent tumor growth *in vivo*. We report here the discovery that the high efficiency of miRNA inactivation in cells by this miRNase is related to its ability to work in a true catalytic mode with multiple turnover, which is increased considerably by recruitment of the non-sequence specific intracellular RNase H.

### MATERIALS AND METHODS

### Peptide-Oligonucleotide Conjugates (POCs)

Oligodeoxyribonucleotides (with and without an aminohexyl linker attached to the 5′-terminal phosphate of the oligonucleotide, and 2′-O-methyl-modification (2′-OMe) (see **Supplementary Table 1**) were synthesized in ICBFM SB RAS (Russia) by Dr. V. Ryabinin and Dr. Maria Meschaninova. The catalytic peptide, acetyl-[(LeuArg)2Gly]2, was attached *via* its C-terminus to the amine group of the aminohexyl linker located at the 5′-terminus of respective oligonucleotides to prepare the miR-21 miRNase and luc-POC conjugates (Patutina et al., 2017a).

### 5**′**-RNA Labeling

miR-21, 5′-UAGCUUAUCAGACUGAUGUUGA-3′, was 5′-end labeled using γ-[32P]-ATP and T4 polynucleotide kinase (Thermo Scientific, USA) as previously described (Silberklang et al., 1979; Mironova et al., 2007).

#### Assay of Ribonuclease Activity and miR-21-miRNase in Multiple Turnover Conditions

Unlabeled miR-21 (10 µM, 25 µM, or 50 µM) and 105 cpm (Cherenkov's counting) of [32P]-miR-21 was incubated at 37°C with either miR-21-miRNase (5 µM) or RNase H (100 U/ml, plus 5 µM of either h-ODN or miR-21-miRNase) in buffer (20 mM Tris-HCl, pH 7.8, 40 mM KCl, 8 mM MgCl2, 1 mM DTT). Reactions were quenched by precipitation of RNA with 2% LiClO4 in acetone, and RNA cleavage products quantified in dried gels as described previously (Patutina et al., 2017a) using Molecular Imager FX and Quantity One software.

#### Nuclease Resistance Assay

Oligonucleotides or conjugates (0.1 µg/µl) were incubated at 37°C in Dulbecco's Modified Eagle Medium (DMEM) (Sigma, USA) with 10% fetal bovine serum (FBS; GE Healthcare, USA). Reactions were quenched in urea (8M) and immediately frozen in liquid nitrogen, prior to analysis of thawed samples in 12% PAAG/8M urea gels, using Tris-Borate-EDTA (TBE) as running buffer, Stains-All (MP Biomedicals, USA) and photographic gel documentation (VilberLourmat, France).

### Cell Lines

Mouse B16 melanoma cells, grown in DMEM/10% FBS/1% antibiotic antimycotic solution (ICN, Germany) and RLS40 lymphosarcoma cells, grown in IMDM/10% FBS/1% antibiotic antimycotic solution, were cultivated at 37°C in a humidified incubator with 5% CO2.

### Transfection of Cells With Oligonucleotides and Conjugates

B16 cells were pre-seeded (2 × 105 per well of 24-well plates) in DMEM/10% FBS a day before transfection. Medium was replaced with serum-free DMEM, prior to incubation for 4 h with h-ODN, 2′-OMe, Inh (hsa-miR-21 5 p inhibitor, Ambion, USA), miR-21-miRNase, or luc-POC, each (0.1 or 1 µM) pre-complexed with Lipofectamine (Thermo Fisher Scientific, USA) in Opti-MEM medium (Thermo Fisher Scientific, USA) according to manufacturer's instructions, then cells were cultivated in fresh culture standard medium for 24–72 h.

### Annexin V-FITC/PI Apoptosis Assay

At 24, 48, and 72 h after transfection, cells were harvested, washed twice with PBS, resuspended in Binding Buffer to a density of 2–5 × 105 /ml, and incubated with annexin V-FITC and propidium iodide (PI) (ab14085, Abcam, UK) in the dark at room temperature for 15 min, prior to flow cytometry analysis (Novocyte CEA Biosciences, USA).

### *In Vitro* Invasiveness Assay

Real-time cell analysis (xCELLigence, ACEABiosciences, USA) used CIM-Plates held at 37°C under an atmosphere of 5% CO2. Top chamber wells were coated with Matrigel (20 µl per well, diluted 1:40 with cold serum-free DMEM) and allowed to polymerize (4 h at 37°C under 5% CO2) prior to cell seeding. Bottom chamber wells were filled with DMEM/10% FBS (160µl each), prior to assembly of the chambers, when serum-free DMEM was added to the top wells (30 µl each) and equilibrated (1 h at 37°C under 5% CO2). At 4 h after transfection, B16 cells (4 × 104 ) were seeded into each top chamber well in serum-free DMEM and pre-incubated for 30 min at room temperature before monitoring (xCELLigence set to collect impedance data, reported as cell index at least once every 30 min).

#### Mice

Male 10–12 week-old CBA/LacSto (hereinafter, CBA) mice were kept in the vivarium of the Institute of Chemical Biology and Fundamental Medicine, SB RAS, with a natural light regime on a standard diet for laboratory animals [GOST (State Standard) R 5025892] in compliance with the international recommendations of the European Convention for the Protection of vertebrate animals used for experimental studies (1997), as well as the rules of laboratory practice in the performance of pre-clinical studies in the Russian State Standards (R 51000.3–96 and 51000.4–96). The experimental protocols were approved by the Committee on the Ethics of Animal Experiments with the Institute of Cytology and Genetics of the Siberian Branch of the Russian Academy of Sciences.

### Tumor Transplantation Assay

RLS40 ascites were taken from CBA mice intraperitoneally injected with tumor cells (2 × 106 in 0.2 ml buffered saline) into their abdominal cavities. RLS40 cells, isolated from ascites fluid by filtration through LSM were divided into seven portions: 1) untreated cells; 2) the cells incubated with Lipofectamine ; the cells transfected with 1 µM of 3) h-ODN, 4) 2′-OMe, 5) Inh, 6) miR-21-miRNase, or 7) luc-POC each. After 4 h of transfection, cell suspensions (0.1 ml, 2 × 106 cells) were intramuscularly inoculated into the right lower limb of CBA mice for solid tumor development. As soon as tumors began to be palpable, the tumor volumes were measured every 2–3 days using calipers. Tumor volumes were calculated as V = (π/6 × length × width × height). Hepatic index was estimated as (liver weight/mouse weight) × 100% and as 5.0% for healthy CBA mice. Tumor doubling time (DT) was estimated as DT = (*t* − *t*0) × ln 2/(ln *V* − ln *V*0), where (*t* − *t*0) indicates the length of time between two measurements of tumor size and *V*0 and *V* denote the tumor volume at two points of the measurement (Ozono et al., 2004).

### STATISTICS

Data were statistically processed using two-way ANOVA taking into account two factors: antisense oligonucleotide and peptide. *Post hoc* testing was completed using Tukey test; *p* < 0.05 was considered to be statistically significant. The statistics package STATISTICA version 12.0 was used for this analysis.

### RESULTS

#### Structure of Nuclease-Resistant miR-21-miRNase

Stability of antisense oligonucleotides (AOSs) against nucleases in serum is essential for *in vivo* application, which typically requires nuclease-resistant chemically modified nucleotides. We avoid this here by design of a nuclease-resistant structure with a hairpin at the 3′-end and the catalytic peptide at the 5′-end. The hairpin oligonucleotide (h-ODN) comprised a 14-mer recognition motif at the 5′-terminus, which is complementary to the miRNA target, and a hairpin structure with a four-member apical loop and 9 nts stem at its 3′-terminus. The catalytic peptide, acetyl- [(LeuArg)2Gly]2, was covalently attached *via* its C-terminus to the 5′-end of the oligonucleotide through an aminohexyl linker (**Figure 1**). The biological and therapeutic properties of miR-21-miRNase was compared with the properties of miR-21-targeted hairpin deoxyribooligonucleotide (h-ODN) and 2′O-methyl-modified oligonucleotide (2′-OMe) unconjugated with the peptide, with a linear 14-mer deoxyribooligonucleotide (ODN) and with the control conjugate (luc-POC), in which the sequence of the hairpin was the same but the complementary part was replaced by the fragment of the luciferase gene with no homology in the mammalian genome (**Supplementary Table 1**). Although the free oligonucleotide (ODN) was chemically unmodified and rapidly degraded in serum, the hairpin ODN resisted dilute serum for a few hours but survived for over 48 h (see **Supplementary Figure 1**), when conjugated to the catalytic peptide within the miRNase (**Supplementary Figure 1**).

#### Induction of Apoptosis and Suppression of Invasiveness of Tumor Cells

By modulating tumorigenesis through the regulation of multiplicity of critical genes suppressing malignant growth, evasion from apoptosis, invasion, and metastasis, miR-21 functions as an oncogene (Melnik, 2015). Knockdown of miR-21 by miR-21 miRNase resulted in specific elevation of the level of the direct mRNA target of miR-21, tumor-suppressor programmed cell death 4 (PDCD4) (Patutina et al., 2017a), and so may also turn down the malignant behavior of tumor cells through induction of apoptosis and inhibition of invasion and metastasis. We studied this in B16 melanoma cells, which are known for maintaining aggressive growth, active migration, and metastasis. As indicated by the Annexin V-FITC/PI assay, miR-21-miRNase increased necrosis of melanoma B16 cells to a limited extent (up to 7%) similar (up to 6.5%) to the sham peptide-oligonucleotide conjugate, luc-POC (for luc-POC sequence see **Supplementary Table 1**) with a sequence of a firefly luciferase gene with no complementarity to the mammalian genome. However, miR-21-miRNase caused a significant increase in the proportion of cells in the apoptotic state with a maximum at 48 h after transfection: the signs of apoptosis were observed in 28% of the cells (cf 15% control), which was six times higher than that in the intact B16 cells (**Figures 2A**, **B**, **Supplementary Figure 2**).

Cells transfected with h-ODN (blue curve), or its methylated derivative 2′-OMe (violet curve) or the sham luc-POC (green curve), migrated at almost the same rate as cells incubated with Lipofectamine (grey curve) (**Figures 2C**, **D**). However, miR-21 miRNase (red curve) strongly inhibited the invasion potential of B16 cells by about 70% relative to control and sham luc-POC (*p*=0.0008). (**Figures 2C**, **D**, time point 40 h). Knockdown of miR-21 by commercial Inh (orange curve) showed only a delayed inhibition of cell invasion of approximately 40% relative to control and only 20% relative to luc-POC.

#### Persistence to Suppress Tumor Growth

To evaluate the ability of miRNase to suppress growth of tumor cells and dissemination in the body, an experiment in mice was conducted. RLS40 lymphosarcoma cells were treated with the miRNase *in vitro* and then intramuscularly transplanted to the right lower limb of CBA mice to form solid tumors with dissemination in the body. miR-21-miRNase efficiency was assessed in a specially collected portion of the treated cells: a 2-fold decrease in miR-21 level by miR-21-miRNase was confirmed by PCR analysis (24 h after transfection), and 1.7- and 2.4-fold increase in protein levels of miR-21 target proteins— PTEN and PDCD4—was shown by Western blot hybridization, respectively, (72 h after transfection) (**Supplementary Figure 3**) which is consistent with previously obtained data (Patutina et al., 2017a). Despite no further treatment of the cells or animals transplanted, the effects of miR-21-miRNase persisted to retard subsequent tumor growth, with a mean tumor volume 16.7 times less than in the control group, showing a 94% tumor volume reduction upon treatment (**Figures 3A**, **B**), which was a statistically reliable difference (*p* value was 10−6 against control). Similar inhibition of tumor growth could only be seen when dosing of the positive controls with Inh was much elevated (1 µM, **Figures 3A, B**) to a 10- to 20-fold higher level than the manufacturer's recommendations (0.05–0.1 μM). Whether the positive control was then specific or caused by a non-specific cytotoxic effect of Inh is unclear. The higher rate of tumor growth in the groups transplanted with RLS40 cells with no treatment (w/t), cells treated with Lipofectamine (LF) alone, Inh (0.1 μM), the methylated h-ODN 2′-OMe (1 μM), and luc-POC did not differ, and h-ODN retarded primary tumor growth at the limit of statistical significance (*p* ~ 0.05) by only about 2-fold (**Figures 3A**, **B**).

Tumor doubling time (DT) was increased to 12.77 ± 0.34 days by miR-21-miRNase, >5-fold greater than of untreated control (w/t) of 2.45 ± 0.49 days (**Figure 3C**). The groups LF, 2′-OMe, luc-POC and h-ODN did not show a statistically significant difference against the w/t, and neither did the positive control of Inh, until a much (10×) elevated level was used to increase

DT to a similar extent (13.90 ± 0.46 days). Relative liver weights remained similar to healthy animals for the miR-21-miRNase and excessive Inh treatments (see **Supplementary Table 2**), even though absence of metastases in the liver may not be expected to persist in tumors seeded from cells treated only before transplant.

### Synergistic Catalytic Recruitment of RNase H

It is desirable that a true artificial nuclease exhibits a reaction catalytic turnover as a result of eventual release of the cleaved RNA fragments followed by attack of the next target miRNA molecules. Using the synthetic 5′-[P32]-labeled miR-21, we evaluated the ability of miRNase to cleave multiple copies of miR-21, when significant excess (up to 10-fold) of miRNA target was used over miR-21-miRNase. Since the recognition motif of miR-21-miRNase was represented by the "naked" (e.g., unmodified) oligodeoxyribonucleotide, it was anticipated that in cellular environment the duplex formed between the miRNase and miR-21 can potentially be recognized by RNase H, which is present in virtually all cells. Therefore, we aimed to evaluate the mutual effect of both RNases on each other (RNase H and miRNase) by comparing their individual activity against miR-21 with their joint action at various excess of miRNA target over miRNase. To achieve this, we ran nine series of the parallel cleavage assays (see **Figures 4A**–**G**). Series A(I), C(IV), and E(VII) (**Figure 4**) measured the efficiency of miRNA cleavage by miRNase alone at the 2-fold, 5-fold, and 10-fold excess of the target RNA with respect to miRNase, respectively. Series A(III), C(VI), and E(IX) evaluated the cleavage efficiency of the RNase H alone towards the miR-21 target present at the same concentrations, while it was hybridized into a heteroduplex with the corresponding oligonucleotide h-ODN lacking the catalytic peptide. Series A(II), C(V), and E(VIII) measured the combined action of miRNase and RNase H present together in the reaction.

We compared the cleavage patterns and kinetics of the miR-21-miRNase with the non-sequence selective RNase H, which is present in virtually all cells. miR-21-miRNase alone cleaved miR-21 at the G3C4, G15A16, G18U19, and G21A22 sites (**Figures 4A**-I, **C-**IV, and **E**-VII), whereas RNase H cuts the target predominantly at the U6A7, C9A10, and G11A12 sites of miR-21 (**Figures 4A**-III, **C**-VI, and **E**-IX). Neither of these cleavage patterns was affected by the concentration of miR-21. The respective activities of miR-21-miRNase and RNase H in combination (**Figures 4A**-II, **C**-V, and **E**-VIII) provided clear evidence that the hetero-complex formed by miRNA

experiments.

and miRNase is also a substrate for RNase H. Target miRNA was effectively cleaved by RNase H when heteroduplexed to the hairpin targeting structure, whether alone (h-ODN) or when conjugated to the peptide (miR-21-miRNase). Moreover, miRNA cleavage occurred at both G-X sites, which are specific for miRNase, and at the sites sensitive to RNase H.

At 2-fold molar excess of miR-21 with respect to h-ODN (**Figure 4A**), the kinetics of miRNA cleavage in a heteroduplex with h-ODN by RNase H alone reached a plateau by 30 min of incubation, but with a total extent of cleavage of only 45% (**Figures 4A**-III, **B**—blue curve). Moreover, with an increase in miRNA concentration to 5- and 10-fold excesses (**Figures 4C**, **E**), the extent of miRNA cleavage by RNase H fell to about 10% and 5%, respectively (**Figures 4C**-VI, **E**-IX; **D** and **F**—blue curves), which presumably reflected the lesser proportions of the miRNA/h-ODN heteroduplex within the overall amount of miRNA present. However, the extent of miRNA cleavage by the miRNase alone was higher and clearly indicative of catalytic turnover. Indeed, the level of cleavage of miR-21 by miRNase alone was similar irrespective of the 2-, 5-, or 10-fold excess of the target over miRNase and progressed towards 65–68% over 24 h (**Figures 4A**-I, **C**-IV, **E**-VII; **B**, **D** and **F**—black curves), and 90% over 72 h (**Supplementary Figure 4A**). Clearly, this miR-21-miRNase structural design was capable of cleaving many copies of the target miRNA, with a greater efficiency than that achieved by a classical antisense pathway *via* RNase H recruitment.

However, as RNase H is ubiquitous in target cells, we also considered whether the simultaneous presence of RNase H affected degradation of miR-21 by this miR-21-miRNase. We discovered that the presence of RNase H led to a considerable increase in the rate and extent of miRNA cleavage, as compared to the individual action of each enzyme alone under identical conditions (**Figure 4**, compare II with I and III, V with IV and VI, and VIII with VII and

FIGURE 3 | Antitumor effect of miR-21-specific miRNase. (A) Retardation of RLS40 growth *in vivo* after treatment of RLS40 cells with antisense oligonucleotides, control conjugate (luc-POC), or miR-21-miRNase. (B) Tumor volume on day 19 after RLS40 transplantation. Mice were injected with RLS40 cells either without any treatment (w/t) or treated with Lipofectamine (LF), commercial miR-21 inhibitor at concentration 0.1 μM (Inh 0.1, recommended) and 1 μM (Inh 1, in accordance to concentration of miRNase used in the study), control conjugate (luc-POC, 1 μM), hairpin 2′OMe oligonucleotide (2′-OMe, 1 μM), hairpin antisense oligonucleotide (h-ODN, 1 μM) or miR-21-specific miRNase (miR-21-miRNase, 1 μM). Data were statistically analyzed using two-way ANOVA with *post hoc* Tukey test. Data are presented as mean ± s.e. *p*-value indicates a statistically reliable difference. (C) Doubling time of RLS40 tumors after treatment with oligonucleotides and conjugates.

FIGURE 4 | Cleavage of 5'-[32P]-labeled miR-21 by miR-21-miRNase and/or RNase H. Autoradiographs of 18% polyacrylamide/8 M urea gel, showing the patterns of cleavage of miR-21 at its 2- (A), 5- (C), and 10-fold (E) molar excess over miR-21-miRNase or h-ODN. Autoradiographs I, IV, and VII represent cleavage of miR-21 by miR-21-miRNase alone, whereas autoradiographs II, V, and VIII show cleavage of miR-21 by a mixture of miR-21-miRNase and RNase H. Autoradiographs III, VI, and IX demonstrate cleavage of the complex formed between miR-21 and h-ODN by RNase H alone (100 U/ml). Duplexes formed by 5′-[32P]-miR-21 (10, 25, and 50 μM) and h-ODN or miRNase (5 μM) were incubated at 37°C for 24–72 h. Lanes Im and T1: imidazole ladder and partial RNA digestion with RNase T1, respectively; "control": RNA was incubated in the absence of oligonucleotide or conjugate and in the presence of RNase H (100 U/ml). Diagrams (B), (D), and (F) show time dependency of cleavage of miR-21 by miR-21-miRNase alone, by a synergetic action of miR-21-miRNase and RNase H, or by RNase H (100 U/ml) alone when miR-21 was in a complex with h-ODN at 2-, 5- and 10-fold molar excess of miR-21 over miR-21-miRNase or h-ODN, respectively. (G) Positions of miR-21 cleavage induced by miR-21-miRNase (red arrows), RNase H in the duplex with h-ODN (black arrows), and by combination of miR-21-miRNase and RNase H. (H) The hypothetical representation of miR-21 cleavage by a combination of miRNase and RNase H.

IX). At a 2-fold excess of miR-21 over miR-21-miRNase, complete (~100%) RNA degradation was observed after 8 h of incubation (**Figures 4A**-II, **B**—red curve, **Supplementary Figure 4B**). With a 5-fold excess, such degradation took 24 h (**Figures 4C**-V, **D** red curve, **Supplementary Figure 4B**), and with a 10-fold excess, RNA degradation was complete within 48 h (**Figures 4E**-VIII, **F**—red curve, **Supplementary Figure 4B**).

By interpolating the half-life (τ/2) of miR-21 degradation, we estimated the catalytic turnover in the absence and presence of RNase H. RNase H elevated cleavage of miR-21 by the miR-21-miRNase by a factor of 14.9 times in the case of 2-fold excess of miR-21, whereas this was less pronounced at 3.0 and 2.4 times for greater excesses of miR-21 at 5-fold and 10-fold, respectively (**Table 1**). In the absence of RNase H, the catalytic turnover of miR-21-miRNase alone was estimated to be around 100 cleaved RNA molecules per molecule of miRNase in 1 h in the cases of 2- and 5-fold excess of miR-21, but reached 382 cleaved RNA molecules per hour at 10-fold miR-21 excess (**Table 1**). The presence of RNase H enhanced the catalytic turnover by factors of 15.6, 7.0, and 4.6 times for 2-fold, 5-fold, and 10-fold excess of miR-21, respectively, with the potential to reach up to 1,800 cleaved miR-21 molecules by one miR-21-miRNase molecule per hour. A possible explanation of such a synergistic effect is that the combined action of the miRNase and RNase H led to formation of very short cleavage fragments of three nucleotides and shorter (**Figures 4A**-II, **4C**-V, **4E**-III) that may rapidly dissociate from the hybridized complex and release the sequencespecific miR-21-miRNase, when it would become available for another cleavage event (see **Figure 4H**). Indeed, the almost complete cleavage of miRNA at 5-fold excess formed short 2 and 3-nucleotide fragments (**Supplementary Figure 5**).

## DISCUSSION

Short functional non-coding microRNAs are implicated in many types of cancer (Frixa et al., 2015; O'Bryan et al., 2017; Zhou et al., 2017). Amongst many miRNAs identified as important inducers of oncogenesis, miR-21 represents an extreme oncomiR, which is strongly involved in tumor onset and progression for many types of malignancies (Huang et al., 2013; Lianidou et al., 2016; Yu et al., 2018). Since miR-21 is abnormally overexpressed in major types of tumors, this oncomiR is the most considered target for anti-miRNA inhibitory studies. However, depending on the origin of tissue and degree of pathological state, the concentration of miRNAs in the cell can vary from several copies to 2–4 μM concentrations, which can be too great to knock down effectively by antisense approaches alone. The most widely used anti-miRNA oligonucleotides, including commercial inhibitors fabricated on the basis of nucleic acids, are usually the perfect complements to miRNA targets, which contain various chemical modifications at the sugar-phosphate backbone to enhance their nuclease resistance and improve binding affinity. However, the chemical substitutions used for generation of such inhibitors are often not compatible with the activity of intracellular RNase H, and their inhibitory effect relies on 1:1 stoichiometry of binding between the oligonucleotide-based inhibitor and miRNA target. Although the use of such oligonucleotide-based inhibitors shows detectable level of miRNA silencing, the absence of the catalytic amplification represents their disadvantage as compared with our biocatalytic agents, capable to degrade irreversibly multiple copies of pathogenic miRNA molecules, which will be particularly crucial to effectively suppress malignant growth. On the basis of our recent discovery of peptide-oligonucleotide conjugates (Patutina et al., 2017a; Staroseletz et al., 2017;

*1)τ/2—a half-life time of miR-21 in the presence of catalysts, the time during which 50% of miRNA molecules is cleaved.*

*2)Rt, reaction turnover—the number of miRNA molecules cleaved by one molecule of miRNase per 1 h. Rt was calculated according to equation Rt = [miR-21] × (number of bonds) × (cleavage, %)/[miRNase].*

Patutina et al., 2018) with desired biocatalytic properties against disease-relevant RNAs, we have designed miRNA-specific aRNase targeted to miR-21. Efficient knockdown of miR-21 by the miR-21-miRNase triggered a broad spectrum of biological responses, leading to the inhibition of pro-survival behavior of tumor cells, including inhibition of tumor cell proliferation by 50% (Patutina et al., 2017a), the initiation of apoptosis in 28% of the tumor cell population, and suppression of cell invasiveness by 60–70%. Furthermore, upon transplantation of the treated tumor cells into mice, the oncosuppressive effect of miR-21-miRNase persisted. Even a single treatment of tumor cells with miR-21 miRNase prior to transplantation led to a 50% reduction in miR-21 level and a drastic (94%) reduction in primary tumor growth in mice. Tumor doubling time increased by more than 5-fold (from 2.45 to 12.77 days) for tumor cells treated with miR-21 miRNase prior to transplantation, as compared to control groups without treatment or treated with a sham (luc-POC) or the hairpin oligonucleotide (h-ODN). The biological effects induced by this miRNase are comparable or superior to those of available miR-21 oligonucleotide inhibitors (Dong et al., 2012; Wagenaar et al., 2015; Wu et al., 2016; Wang et al., 2018; Zhou et al., 2018).

The design developed for this miRNase: i) enhanced the hybridization efficiency towards the miRNA target, probably due to the additional stacking interactions provided by the hairpin structure (Sunami et al., 2004; Patutina et al., 2017b); ii) bound only mature forms of miRNA; and iii) protected the oligonucleotide from nuclease degradation. It is well known that high nuclease susceptibility of natural oligodeoxyribonucleotides leads to short half-life and reduced antisense functionality *in vivo*, typically requiring the use of chemically modified oligonucleotides to avoid this rapid degradation. However, the rational design of nuclease resistance into these therapeutic miRNases was most likely provided by the hairpin targeting structure and the catalytic peptide location. The h-ODN with a hairpin at the 3′-end remained intact for a few hours but, with the catalytic peptide conjugated at the 5′-end of the hairpin oligonucleotide, persisted for more than 24 h. This nuclease resistance of the miR-21-miRNase undoubtedly contributed to the longer and stronger biological effect observed *in vitro* and *in vivo*. The nuclease resistance of this miRNase is not inferior to the stability of potent phosphorothioate DNA/LNA mixmer, which begins to degrade by 24 h of similar incubation (Lennox and Behlke, 2010). The obtained results correlate with the data reported earlier, where it was shown that 3′-end structures can improve resistance to snake venom phosphodiesterase, DNA polymerase I, and FBS and significantly increase functional potency for target RNA knockdown (Tang et al., 1993; Hosono et al., 1995). The presence of flanking duplexes also improves the ability of anti-miRNA oligonucleotides to invade RISC and serve functional enhancement (Vermeulen et al., 2007; Lennox and Behlke, 2011).

Clear evidence of the catalytic nature of miRNA cleavage by this miRNase was found when the miR-21 was present at 2-, 5-. or 10-fold excess over the miRNase. In contrast, RNase H alone did not exhibit any catalytic turnover, as the extent of miR-21 cleavage by this ribonuclease simply correlated with the proportion of the miR-21/h-POC hetero-complex within the overall amount of miRNA present, which is the expected result for a conventional antisense-mediated pathway involving RNase H recruitment. Interestingly, neither introduction of the hairpin nor the presence of the peptide within the structure of the miR-21-miRNase affected the ability of RNase H to recognize and cleave miR-21 hybridized with the oligonucleotide moiety of the miR-21-miRNase.

Particularly exciting though was the discovery of the synergistic effect from the joint action of this miRNase and RNase H towards miR-21. In isolation, both RNases cleaved totally different regions of miR-21, which can be easily identified from the PAAG analysis (**Figure 4**). However, when miR-21 formed a complementary duplex with h-ODN oligonucleotide, RNase H recognized this hetero-complex and cleaved miRNA at the "seed region," located close to the 5′-end (bases 2–8), which is known to be a "canonical" determinant in miRNA function (Lai, 2002). In contrast, miR-21 miRNase cleaved the 3′-region of the miR-21 molecule (bases 13–18), which represents a "3′-compensatory" or "beneficial 3′-paring" site in the miRNA-mRNA target recognition and plays a crucial role for miRNA specificity and functioning (Brennecke et al., 2005; Grimson et al., 2007; Wang et al., 2009; Robertson et al., 2010). However, when both ribonucleases were present (which appears likely *in vivo* in the intracellular environment), we observed a significant increase of the rate of miRNA cleavage. Earlier, using miRNA-targeted conjugates of a different structure, we have shown that the presence of RNase H can significantly enhance its activity and increase the rate of target RNA cleavage, without any positive effect on RNase H activity (Patutina et al., 2018). In this study, we discovered a different mechanism of synergy when both RNases showed mutual enhancement of their cleavage activity in the presence of each other, which was considerably higher than any simple additive effect expected from independent enzymatic action. The observed synergistic effect from this combined attack on miRNA molecules could be attributed to multiple cuts within miR-21 induced by these ribonucleases, thus leading to a formation of short cleavage products. This may trigger a rapid collapse of the hybridized complex and a release of both miRNase and RNase H, when each could then initiate a new attack on another miR-21 molecule (**Figure 4H**). This mechanism may well explain the high level of inactivation of oncogenic miRNA and the persistence of diverse suppression of malignant behavior of tumor cells upon treatment with this miR-21-miRNase.

We conclude that high therapeutic efficacy of peptideoligonucleotide conjugates is likely to arise from the following: 1) high resistance to nuclease degradation from structural design, 2) efficient cleavage of miRNA-target molecules in a multiple turnover or true catalytic manner, and 3) by synergistically harnessing intracellular RNases such as RNase H. Over the past few decades, many different artificial ribonucleases have been reported, with some cleaving RNA *in vitro* with high efficiency and selectivity. However, to the best of our knowledge, we report here the first example of metal-independent, RNAspecific peptide-oligonucleotide conjugates operating under physiologically relevant conditions within tumor cells, which maintained miRNA knockdown in tumors subsequently formed from cells transplanted into mice, without further treatment. These findings elevate these sequence-specific aRNases to worthy rivals to antisense and siRNA technologies for the development of new RNA targeting therapeutics.

The key discoveries of this research provide a solid experimental ground for facilitating a paradigm shift in the development of new therapeutic interventions to ultimately offer effective, safe and cost-efficient treatment of unresectable or metastatic tumors, which currently relies on the cytotoxic effect of chemotherapies or radiotherapy and inevitably leads to a severe toxicity in humans. A fundamental change in treatment from smallmolecule anti-cancer drugs, which often suffer from poor target selectivity, to highly selective RNA-targeting therapies provides a possibility to overcome such undesirable side effects arising from cytotoxic drug cocktails of combination regimens currently used in oncology. Our miRNA-selective chemical ribonucleases offer unprecedented opportunity to selectively knock down many copies of oncogenic miRNA in cells by irreversible cleavage in a truly catalytic manner with multiple turnovers, which is reinforced even further by recruitment of intracellular RNase H. This will ultimately boost potency, reduce dosage, and decrease cost of treatment. In the future, miRNases can be used both as a monotherapy and as an adjuvant therapy in combination with surgical treatment, radiotherapy, and chemotherapy. In the latter case, miRNases may allow to significantly decrease the dosage of cytostatics thereby reducing toxicity. Application of miRNases is a promising and rapidly evolving area of antisense technology, and its suitable combination with chemotherapeutics may represent highly efficient approaches to treating oncopathologies and other miRNA-associated diseases in humans.

#### DATA AVAILABILITY

All datasets generated for this study are included in the manuscript/supplementary files.

### ETHICS STATEMENT

This study was carried out in accordance with the international recommendations of the European Convention

#### REFERENCES

for the Protection of vertebrate animals used for experimental studies (1997), as well as the rules of laboratory practice in the performance of pre-clinical studies in the Russian State Standards (R 51000.3–96 and 51000.4–96), the Committee on the Ethics of Animal Experiments with the Institute of Cytology and Genetics of the Siberian Branch of the Russian Academy of Sciences. The protocol was approved by the Committee on the Ethics of Animal Experiments with the Institute of Cytology and Genetics of the Siberian Branch of the Russian Academy of Sciences.

#### AUTHOR CONTRIBUTIONS

MZ, EB, and OP conceptualized and designed the study. VV supervised the research. OP, SM, NM, AS, and EB performed the experiments and analyzed the data. OP, EB, DC, and MZ interpreted the data, wrote, edited, and revised the manuscript. All authors approved the final version of the manuscript.

#### FUNDING

This work was funded by Russian Science Foundation (Grants No. 14-44-00068 and No. 19-14-00250) and by Russian State funded budget project of ICBFM SB RAS # АААА-А17-117020210024-8, BBSRC (Grant No. BB/K012622/1), and EPSRC (Grant No. EP/ E003400/1).

#### ACKNOWLEDGMENTS

The authors are grateful to Mrs. Albina V. Vladimirova (ICBFM SB RAS) for cell maintenance.

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fphar.2019.00879/ full#supplementary-material

distant metastasis and survival in kidney renal clear cell carcinoma. *J. Cancer* 9, 3651–3659. doi: 10.7150/jca.27117

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2019 Patutina, Miroshnichenko, Mironova, Sen'kova, Bichenkova, Clarke, Vlassov and Zenkova. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# RNases Disrupt the Adaptive Potential of Malignant Cells: Perspectives for Therapy

*Vladimir Alexandrovich Mitkevich\*, Irina Yu Petrushanko and Alexander Alexander Makarov*

*Engelhardt Institute of Molecular Biology, Russian Academy of Sciences, Moscow, Russia*

Exogenous RNases are selectively toxic to tumor cells. The reasons for this selectivity are not quite clear and should be searched for in the properties that distinguish malignant from normal cells. During onco-transformation, cells acquire properties allowing them to adapt to the altered microenvironment, such as resistance to hypoxia, changes in intracellular pH, disruption of ion transport, reduced adhesion and increased mobility, and production of specific exosomes. These adaptation mechanisms distinguish malignant cells from normal ones and give them a competitive advantage, ensuring survival and spread in the organism. Here, we analyze if the directed cytotoxic effect of exogenous RNases is linked to the disruption of the adaptive potential of tumor cells and how it can be used in anticancer therapy.

#### *Edited by:*

*Jean-Michel Fustin, Kyoto University, Japan*

#### *Reviewed by:*

*Pavel Ivanov, Brigham and Women's Hospital, United States David Pulido-Gomez, University of Oxford, United Kingdom*

### *\*Correspondence:*

 *Vladimir Alexandrovich Mitkevich<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Translational Pharmacology, a section of the journal Frontiers in Pharmacology*

*Received: 10 April 2019 Accepted: 22 July 2019 Published: 16 August 2019*

#### *Citation:*

*Mitkevich VA, Petrushanko IY and Makarov AA (2019) RNases Disrupt the Adaptive Potential of Malignant Cells: Perspectives for Therapy. Front. Pharmacol. 10:922. doi: 10.3389/fphar.2019.00922*

Keywords: RNase, malignant cell, adaptation potential, external environment, redox state

### INTRODUCTION

Enzymes that hydrolyze RNA, ribonucleases (RNases), have several biological effects on cells. Stimulation of the growth of blood vessels, toxicity toward tumor cells, and antiviral activity are important therapeutic properties of RNases.  However, the mechanism of their antitumor activity remains largely unclear. RNases are thought to exhibit cytotoxic activity by cleaving cellular RNA. The most accessible for them are RNAs that are not associated with proteins, i.e., transport and non-coding miRNAs. Thus, angiogenin cuts tRNA molecules inside anticodon loops, producing "stress-induced" RNA fragments, which leads to inhibition of translation initiation (Ivanov et al., 2011). The cytotoxicity of onconase, RNAse from *Rana pipiens*, is also associated with the degradation of tRNA in the cytosol of tumor cells with subsequent inhibition of protein synthesis and apoptosis (Saxena et al., 2002). Сytotoxic RNAse α-sarcin from *Aspergillus oryzae* specifically cleaves one bond in 28S rRNA, causing inhibition of translation and subsequent cell death (Olmo et al., 2001). However, numerous evidences indicate that inhibition of protein synthesis is not the only cause of apoptosis induced by RNases (Qiao et al., 2012). In particular, it has been shown that binase, RNase from *Bacillus pumilus*, directly interacts with K-RAS oncoprotein, disrupting its functioning (Ilinskaya et al., 2016). No correlation was found between the decrease in the level of RNA and the toxic effect of RNases. Thus, in the precursors of myeloid cells, binase reduced the level of RNA by 20% with 67% cell viability, and in the human kidney epithelium cells, RNA levels were halved while 85% of the cells remained viable (Ilinskaya et al., 2008). In the cells of acute myeloid leukemia Kasumi-1, which are extremely sensitive to binase, the total RNA level did not change even when the viability decreased by 95% (Mitkevich et al., 2011). Onconase caused apoptosis of mitogen-stimulated lymphocytes, without reducing the level of cellular RNA (Ardelt et al., 2003).

Another argument in favor of the fact that the degradation of available RNA is not the only cause of the cytotoxicity of RNases is that they selectively eliminate certain malignant cells. It is known that the malignancy of cells occurs as a result of a change in the processes of differentiation and control of division, which may be due to the expression of oncogenes. At the same time, several similar cancerous cell lines may respond differently to RNases (Altomare et al., 2010).  Apparently, sensitivity to RNases occurs due to the activation and/or inhibition of certain signaling pathways and changes in the properties of onco-transformed cells. It is also obvious that the rate of cell division is not a decisive criterion for the toxic action of RNases. In accordance with our hypothesis, based on the study of the molecular mechanisms of the cytotoxic effect of exogenous RNases in recent years (Mitkevich et al., 2011; Mironova et al., 2013; Mitkevich et al., 2014; Mitkevich et al., 2015; Mitkevich et al., 2017), the reason for the selective toxicity of RNase to cancer cells should be sought in the properties of tumor cells that enable them to constantly adapt to the extracellular environment. Unlike normal cells that exist in conditions of permanent tissue homeostasis, actively proliferating malignant cells affect the composition and properties of their microenvironment and must continuously adapt to changing environmental conditions. The properties of malignant cells that ensure their survival, proliferation, and distribution include altered redox status, resistance to hypoxia, changes in the functioning of ion transport systems, increased intracellular pH, reduced adhesion, and production of specific exosomes. Despite an intensive study of the toxic properties of RNases from different organisms, information about their influence on the above properties of tumor cells is rather scarce. Here, we summarize the knowledge accumulated to date about influence of RNases on adaptive properties of malignant cells and consider the directions of future investigations for RNases application in the treatment of tumors.

#### RNASES AND INTRACELLULAR REDOX STATUS

Cancer cells possess a redox potential different from normal cells due to their accelerated metabolism. They demand high levels of reactive oxygen species (ROS) to maintain a high proliferation rate (Sosa et al., 2013). The thiol redox status, which depends on the ratio of reduced (GSH) and oxidized (GSSG) glutathione (Gonzalez-Dosal et al., 2011; Wang et al., 2013), is also altered in cancer cells. Under normal conditions, the GSH level in cells (1–5 mM) is 100-fold higher than GSSG. Under oxidative stress, this ratio can be reduced to 1 (Allen and Mieyal, 2012). However, many tumors show elevated levels of GSH emphasizing the link between the deregulations of GSH homeostasis and cancer (Tew and Townsend, 2011). Changes in the redox status lead to changes in the functioning of redox-sensitive proteins (kinases, transcription factors, ionic transporters, etc.) due to their redox modification (Gao and Schottker, 2017). In particular, the shift of intracellular redox conditions to the oxidized state induces protein glutathionylation that protects the thiol groups of proteins from irreversible oxidation and changes their function (Mieyal et al., 2008; Miller and Mieyal, 2015). In the case of viruses that provoke oncogenesis, a change in the redox status is necessary to "tweak" the cell to fit their needs. Thus, human papillomavirus (HPV) suppresses antioxidant systems in the cell for the functioning of redox-sensitive viral oncoproteins, E6 and E7, which bind to the tumor suppressors p53 and retinoblastoma protein, respectively, suppressing their activity, which stimulates uncontrolled proliferation and prevents apoptosis. The return of cells to normal redox status will reduce the activity of such oncoproteins. Redox active drugs, which are based on changing the cell's redox status, have already proven their effectiveness in the treatment of certain types of cancer including promyelocytic leukemia, esophageal, ovarian, non-small cell lung, colon, and breast cancer (Wondrak, 2009; Tew and Townsend, 2011). It has been established that RNases lead to a decrease in the level of ROS in tumor cells. So, onconase caused a decrease in ROS in cells of acute lymphoblastic leukemia, Jurkat, and several fibroblast cell lines (Ardelt et al., 2007). Binase caused a decrease in ROS level in Kasumi-1 cells, B16 mouse melanoma, and Jurkat cells (Mironova et al., 2013; Mitkevich et al., 2013; Burnysheva et al., 2016). RNases, by reducing ROS in tumor cells, alter the redox modifications of key proteins (such as NF-kB, p53, etc.), which, in turn, suppress the resistance of cancer cells to apoptosis. Thus, the "return" of the redox status of cancer cells by RNases to the values characteristic of normal cells lead to a decrease in their resistance to death and uncontrolled division by normalization of the redox-sensitive cellular systems.

It should be mentioned, that one of the key player in redox regulation of cells is cytosolic RNase inhibitor (RI) (Dickson et al., 2005). It binds to and inhibits extracellular RNases. It was postulated that only RNases that evade RI can kill cells. For example, RI regulates angiogenesis through very strong binding to angiogenin, which is a cytoprotective RNase. However, in the absence of RI, the angiogenin converts into a cytotoxic molecule (Thomas et al., 2018). Unlike mammalian RNases, RI does not inhibit the ribonuclease activity of cytotoxic bacterial RNases (Sevcik et al., 2002), but the absence of direct interaction between them has not been shown. RI is very sensitive to oxidative stress and is easily inactivated, while losing its ability to inhibit exogenous RNases (Dickson et al., 2005). RI inactivation can work as mechanism to switch on the RNase-mediated degradation of cellular RNA under stress conditions (Lomax et al., 2014). On the other hand, RI protects cells from oxidative stress, and its deprivation leads to decrease of GSH level as well as increase of oxidant-induced DNA damage (Monti et al., 2007). Decrease of RI activity in tumor cells induces their proliferation, increases migration and invasion, and reduces their adhesion (Chen et al., 2011). Upregulation of RI activity, on the contrary, causes the death of tumor cells (Tang et al., 2019). One can suggest that cytotoxic RNase influence on RI status by several ways. RNase reduction of ROS level, increased in tumor cells, protects RI from inactivation; exogenous RNase in cells can upregulate RI, which will reduce oxidative stress; interaction of RI with RNase may not lead to inactivation of the latter; however, it may change the properties of the RI, including protection against oxidation. These points should be clarified in future studies.

### RNASES AND HYPOXIA

Uncontrolled division of tumor cells changes their normal microenvironment. In particular, due to the lack of blood supply, cells at a certain stage of tumor development are in a state of hypoxia. Hypoxia serves as an additional risk factor that accelerates cell malignancy. Under hypoxia and/or altered redox status of cancer cells, the transcriptional factor hypoxia inducible factor 1 (HIF1) stabilizes in them, which makes the cells resistant to oxygen deficiency. Hypoxia is a characteristic marker for the development of tumors and has a significant impact on the course of chemotherapy, often negative (Carreau et al., 2011). It can also affect the expression level of oncogenes. In particular, during hypoxia, miR29b synthesis is suppressed, which normally suppresses the synthesis of KIT oncoprotein (Liu et al., 2010) and, consequently, the level of KIT in cells increases. Activation of KIT leads to an increase in the level of HIF1 (Zhang et al., 2011). It was shown that co-expression of AML1-ETO and HIF1 oncogenes in myelogenous leukemia cells leads to a higher rate of cell proliferation *in vitro* and to a more severe course of leukemia in mice (Gao et al., 2015). By itself, stabilization of HIF1 under conditions of chronic hypoxia is a risk factor for the spread of a tumor.

The work of endogenous RNases designed to suppress the development of neoplasia is impaired under hypoxic conditions. Nature has provided a mechanism for the destruction of cells with stabilized HIF1 by increasing the level of endogenous RNase T2 (Uccella et al., 2018). It inhibits angiogenesis and induces apoptosis of malignant cells. However, in aggressive cancers, the inhibition of cell growth by RNase T2 stops working. Reduced expression of DICER, the enzyme involved in microRNA processing, is frequently observed in cancer and is associated with poor clinical outcome in various malignancies. DICER expression is suppressed by hypoxia through an epigenetic mechanism that involves inhibition of oxygen-dependent H3K27me3 demethylases KDM6A/B and results in silencing of the DICER promoter. Subsequently, reduced miRNA processing leads to de-repression of the miR-200 target ZEB1, stimulates the epithelial to mesenchymal transition, and ultimately results in the acquisition of stem cell phenotypes in human mammary epithelial cells (Van den Beucken et al., 2014). It was found that miR-630, which is upregulated under hypoxic conditions, targets and downregulates DICER expression. In an orthotopic mouse model of ovarian cancer, delivery of miR-630 using 1,2-dioleoyl-sn-glycero-3-phosphocholine nanoliposomes resulted in increased tumor growth and metastasis and decreased DICER expression (Rupaimoole et al., 2016). Exogenous RNase, resistant to inactivation under conditions of altered redox status during hypoxia, can compensate for the loss of endogenous RNase function. Binase does not contain cysteine and methionine residues and, accordingly, is insensitive to changes of redox conditions. We tested the effect of binase on Kasumi-1 cells, and cervical cancer SiHa cells growing at different oxygen contents. A decrease in [O2] from 21 to 5 and 1% resulted in an almost two-fold increase in the proportion of apoptotic cells in the Kasumi-1 and SiHa cells treated by binase. The increased sensitivity of cancer cells to the effect of binase under decreased oxygen level is associated with a change in the expression of oncogenes and the activation of processes mediated by oncogenic proteins. This suggests that the response of malignant cells to RNases during tumor development may be enhanced by disrupting their adaptation to low oxygen conditions.

### RNASES AND PH

Disruption of oxygen supply leads to aerobic glycolysis in cancer cells (Warburg effect). The excess of protons produced during glycolysis, by the Na+/H+ exchanger is transferred to the extracellular environment. Change in pH is one of the markers of cancer cells (Webb et al., 2011). Malignant cells have a "reversed" pH gradient with a constantly elevated intracellular pH that is higher than the extracellular pH (Webb et al., 2011). The increase in intracellular pH leads to the induction of cell proliferation, increases their resistance to apoptosis, and metabolically adapts the cells to oxygen deficiency. It has been suggested that a decrease in the intracellular pH of malignant cells will lead to antitumor effects (Slingerland et al., 2013). We obtained preliminary data showing that the effect of binase (0.8 µM) on Kasumi-1 cells under 20%, 5%, 1%, and 0.2% [O2] leads to a decrease in the intracellular pH value by 0.2–0.5 units. Since the normal functioning of tumor cells is associated with high values of intracellular pH, the decrease of this parameter under the action of RNase should disrupt the intracellular regulation and reduce their adaptive potential.

### RNASES AND ADHESION

To adapt to the extracellular environment, cancer cells re-arrange their plasma membranes to sustain proliferation, avoid apoptosis, and resist anticancer drugs. This leads to changes in the cell deformability, which is important for invasiveness, membrane stiffness, and receptor function causing disruption of adhesion and intercellular signaling (Bernardes and Fialho, 2018). Membrane of normal cells is characterized by a lipid asymmetry between the inner and outer leaflets. A de-regulation of this asymmetry is often encountered in cancerous cells where phosphatidylserine is often exposed in the outer membrane resulting in a negative surface charge, leading to disruption of cell adhesion and promotion of metastasis and tumor invasion (Bernardes and Fialho, 2018). It has been shown that an increase in intracellular pH is necessary for the directed migration of malignant cells (Webb et al., 2011). During oncogenic transformation of cells, there is also a change in the structure and expression of the glycoproteins and glycolipids in the cell membrane, which form the so-called adhesive molecules (Hart et al., 1991). As a result, the membrane of tumor cells contains more acid glycoproteins and phospholipids than the membrane of normal cells. This adaptation marker is one of the reasons for the selective elimination of tumor cells by RNases. Indeed, the majority of known cytotoxic RNases are basic proteins, and cationization of RNases is considered to be an effective strategy for strengthening their antitumor properties (Mitkevich et al., 2014). The proliferation of tumor cells may be associated with disruption of endogenous RNases. It has been demonstrated that endogenous RNase L inhibits cell attachment and cell spreading (Dayal et al., 2017). Prostate cancer cells depleted of RNase L show greater migration in wound healing and trans-well migration assays in response to fibronectin and serum. Over-expression of RNase L suppressed cell migration compared to both endogenous levels and knockdown cells while activation of RNase L, which requires RNase L dimerization, which inhibited cell migration. This was attributed to the destabilization of the miR-regulated transcriptome by RNase L (Rath et al., 2015). However, it was later shown that the effect of RNase L on cell migration is mediated, in part, by protein–protein interactions and does not require enzymatic activity (Dayal et al., 2017). We have previously established that binase reduces metastasis in animals (Mironova et al., 2013). This allows suggesting that binase can reduce invasion and enhances the adhesion of tumor cells. One can expect that exogenous RNases are able to influence the adhesion and motility of tumor cells, preventing the spread of the tumor. It should be borne in mind that such effects on tumor cells may have upregulation of RI, so it is absolutely necessary to verify the role of cytotoxic RNases in regulating RI activity.

### RNASES AND ION TRANSPORT

Change in the functioning of ion transport systems plays a major role in the adaptability of malignant cells. The increase in intracellular pH in tumor cells is associated with changes in the activity of ionic transporters. Neoplasms induced by oncogenes or carcinogens have an abnormally high Na+ content, caused by membrane depolarization and the opening of potential-dependent Na+ channels (Lobikin et al., 2012). Membrane depolarization serves as a marker for carcinogenesis and suggests that the development of neoplastic changes can be delayed by increasing the expression of ion channels, which leads to hyperpolarization of membranes for example, K+ and Cl channels. Some pharmacological antitumor agents act precisely by changing the activity of ion channels (Lobikin et al., 2012; Peigneur et al., 2012; Schweikart et al., 2013). We have found that cytotoxic bacterial RNases, binase, and 5K RNase Sa (highly cationic mutant of RNase Sa from *Streptomyces aureofaciens*) can change the functional activity of calcium-activated potassium channels artificially introduced into embryonic kidney cells (Ilinskaya et al., 2004; Ilinskaya et al., 2008). Binase leads to an increase in the level of Ca2+ (Mitkevich et al., 2013), which may be due to inhibition of the Na,K-ATPase, leading to the activation of the Na+/Ca2+ exchanger. Na,K-ATPase is the main ion transport system in mammalian cells. It creates a transmembrane gradient of K+ and Na+ ions in cells. This pump plays an important role in the regulation of cell volume, transmembrane potential, intracellular pH, and Ca2+ level through Na+/H+ and Na+/Ca2+ exchangers (Slingerland et al., 2013). In tumor cells, the activity of Na,K-ATPase is increased, which may be due to the need to remove Na+, which accumulates as a result of intensive activity of the Na+/H+ exchanger. In addition, maintaining the level of intracellular K+ with Na,K-ATPase is also necessary to suppress apoptosis. Our preliminary experiments showed that, in Kasumi-1 cells, under the action of binase (0.8 µM, 24 h), the hydrolytic activity of Na,K-ATPase was reduced by 80% (**Figure 1A**). This is due to both a 25% decrease in the level of the enzyme in cells (**Figure 1B**) and an increase in its glutathionylation level by 50% (**Figure 1C**, **D** ). It was shown that glutathionylation led to inactivation of Na,K-ATPase and associated with a change in the cell redox status (Petrushanko et al., 2012). The decrease in the activity of Na,K-ATPase leads to a violation of ionic homeostasis and precedes the manifestation of the apoptogenic effects of binase. Obviously, this is one of the components of the mechanism of the cytotoxic effect of binase on onco-transformed cells. Thus, the effect of exogenous RNases on ion transport systems may be another reason for the selective cytotoxicity of RNases on cancer cells.

### RNASES AND VESICULAR TRANSPORT

Exosomal vesicles secreted by tumor cell play an active role in oncogenesis and metastasis (Kim et al., 2017). They communicate between tumor cells and their microenvironment. Exosomes transporting miRNAs and proteins can cause neoplastic transformation and aid tumor development (Zhang et al., 2015). miRNAs absorbed by the recipient cell from exosomes can act as a regulator of gene expression; there is growing evidence that they are involved in the onco-transformation of cells (Kim et al., 2017). RNases bound to negatively charged molecules on the surface of cells penetrate them by endocytosis. It has been established that RNase A and BS-RNase bind actin *in vitro* and induce its polymerization (Simm et al., 1987). Since the process of polymerization/depolymerization of actin governs its dynamic properties during endocytosis, it can be assumed that RNases can influence the endocytosis of tumor cells, changing their interaction with the microenvironment.

We did not find information on how RNases influence production and composition of the exosomes of malignant cells. However, it can be assumed that since RNases change the properties of tumor cells, affecting ionic homeostasis and redox status, the number and composition of the exosomes produced by the cells will also change. An indirect confirmation of our assumptions may be the data from (Mironova et al., 2013). The authors found that RNase A reduces the level of miRNA in blood stream of mice with Luis lung carcinoma. This decrease is not related to the level of miRNA in tumor cells. MiRNA degradation by RNase A did not observed *in vitro* conditions. Given that part of extracellular miRNAs in the blood is enclosed in exosomes, it can be assumed that the decrease of miRNA level is associated with a decrease in the production of exosomes by cancer cells after treatment by RNase A. Probably, RNases affect the number and properties of RNA molecules entering into exosomes. In this case, the cancer cell will lose "communication channels" with the microenvironment, which will inevitably reduce its adaptive potential and reduce the risk of tumor spread.

### IMMUNE-MODULATORY EFFECTS OF RNASES

One of the main factors in the survival of tumor cells in the organism is their ability to circumvent immune surveillance. Tumor cells may escape from immune control and proliferate in an unrestricted manner (Mittal et al., 2014). This escape can be mediated through various mechanisms, such as reduced immune recognition, increased resistance to attack by immune cells, or the development of an immunosuppressive tumor microenvironment (Mittal et al., 2014). Human

S-glutathionylated (GSS-α1/α1) form of the protein normalized to its total amount. The levels of S-glutathionylated Na,K-ATPase α1-subunit and total α1-subunit were estimated using immunoblotting as described in (Mitkevich et al., 2016). Mouse monoclonal anti-glutathione antibody MAB5310 (Millipore) and mouse monoclonal anti-Na,K-ATPase α1 antibody clone C464-6 (Millipore) were applied to detect glutathionylated proteins and total amount of α1-subunit correspondingly. Mouse monoclonal antibody AM4302 (Ambion) was used for β-actin detection. After staining with horseradish peroxidase–conjugated secondary antibodies, membrane was stained using a commercial kit SuperSignal West Femto Maximum Sensitivity Substrate (Thermo Scientific), and chemiluminescence was detected using Bio-Rad ChemiDoc MP instrument. Densitometric analysis was performed by Image Lab (Bio-Rad) program, and the results were represented as ratio of glutathionylated α1- subunit to total α1-subunit band intensity ([GSS- α1]/total α1) or as ratio of α1-subunit to β-actin band intensity (α1/β-actin). (D) The original immunoblotting readouts. Data are mean values for three independent experiments with triplicates ± standard diviation. The differences among the groups were analyzed by Student's t-test, and p < 0.05 was considered statistically significant. Statistica 7 software was used for analysis. \*p < 0.005.

extracellular RNases are mainly expressed in innate cells and display a variety of immune modulation activities (Lu et al., 2018). They can participate in host immune responses, working as alarmins and safeguard molecules against infection and inflammation. Secretion of RNase in the focus of inflammation сontributes to tissue repair and remodeling. RNases also function as cytokines and chemokines, displaying anti-inflammatory activities and inducing chemoattraction of innate cells (Lu et al., 2018). Activity of host RNases is often lowered in tumor tissue (Shlyakhovenko, 2009) and significantly reduced in serum of patients with different types of cancer (Huang et al., 2014). It turned out that exogenous RNases also have an immunomodulatory effect (Lu et al., 2018) and can be used to replenish the pool of extracellular RNases and modulate the immune response. For example, macrophage immune regulation by binase can trigger the host cell antitumor response (Makeeva et al., 2017). Binase also showed its effectiveness in restoring interferon sensitivity (IFN) of SiHa cervical cancer cells, in which the IFN response was initially suppressed by HPV (Mitkevich et al., 2017).

### DISRUPTION OF TUMOR CELL ADAPTABILITY BY RNASES AS A POTENTIAL CANCER THERAPEUTIC TARGET

From the presented data, it is evident that the influence of exogenous RNases on the adaptation of malignant cells to the external environment has not been properly explored. Such adaptation mechanisms give malignant cells competitive advantages, ensuring their survival and distribution in the body. Influencing these mechanisms with the help of exogenous RNases will allow the weakening of the adaptation potential of tumor cells, inducing their death and making them more susceptible to therapeutic agents (**Figure 2**). A comprehensive

FIGURE 2 | Disruption of the adaptation mechanisms of cancer cells by RNase. Upper panel: cancer cell adapts to the environment (decreased pO2 level) and changes it for itself, thus avoiding the immune system cells and drugs. The properties of malignant cell that ensure its survival, proliferation, and spread include altered redox status (increased reactive oxygen species [ROS]), resistance to hypoxia (hypoxia inducible factor [HIF1] increased and stabilized), changes in the functioning of ion transport systems (dysregulation of N+,K+ and H+,K+ pumps and exchangers), increased intracellular pH value, reduced adhesion, disruption of endogenous RNases (endoRNase), and production of specific exosomes (vesicles) containing specific cancer miRNA. Lower panel: exogenous RNases (exoRNase, green stars) act on a whole range of adaptive properties of tumor cells—decrease level of HIF1, ROS, and pH value; reduce activity of ion transporters; suppress exosomes production and degrade miRNA; and stabilize cell adhesion. These make tumor cells available for elimination by the immune system and reduce their drug resistance.

### REFERENCES

study of the adaptive properties of cells that are influenced by RNases will determine the mechanisms of their selective cytotoxicity to malignant cells and establish conditions affecting the effectiveness of their antitumor activity. This will make it possible to effectively use RNases in combination with other therapeutic agents to which the tumor cells are initially resistant. Drug resistance of tumor cells is largely related to their ability to adapt to the environment. Reducing the adaptive capacity of neoplastic cells with RNases can be an effective strategy for overcoming drug resistance. A similar approach was demonstrated by us using the example of SiHa cells, the oncotransformation of which is due to HPV. Viral oncoproteins suppress the IFN response in cells, as a result of which they become resistant to interferon therapy. Binase restores IFN signaling, enhances IFN sensitivity and apoptosis in SiHa cells (Mitkevich et al., 2017). The advantage of binase and, probably, other exogenous RNases is that under conditions of low oxygen content in the environment its antineoplastic properties are increased, which accentuates the therapeutic potential of RNase for chemo-resistant tumors. Since, RNases act on a whole range of adaptive properties of tumor cells and retain their activity even in a changing micro-environment (with altered redox status, pH, oxygen, etc.), they are promising molecules for the treatment of various types of tumors. The combination of RNases with other types of anticancer drugs can help solve the problem of drug resistance of tumor cells.

### DATA AVAILABILITY

All datasets generated for this study are included in the manuscript and the supplementary files.

### AUTHOR CONTRIBUTIONS

All authors listed have made substantial, direct and intellectual contribution to the work, and approved it for publication.

### FUNDING

The study was supported by Russian Foundation for Basic Research (Grant #17-00-00061).

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Shlyakhovenko, V. A. (2009). Ribonucleases in tumor growth. *Exp. Oncol.* 31, 127–133.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2019 Mitkevich, Petrushanko and Makarov. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Characterization of the *Puumala orthohantavirus* Strains in the Northwestern Region of the Republic of Tatarstan in Relation to the Clinical Manifestations in Hemorrhagic Fever With Renal Syndrome Patients

*Yuriy N. Davidyuk1, Emmanuel Kabwe1, Venera G. Shakirova2, Ekaterina V. Martynova1, Ruzilya K. Ismagilova3, Ilsiyar M. Khaertynova2, Svetlana F. Khaiboullina1,4, Albert A. Rizvanov1 and Sergey P. Morzunov5\**

*1 OpenLab Gene and Cell Technologies, Institute of Fundamental Medicine and Biology, Kazan Federal University, Kazan, Russia, 2 Department of Infectious Diseases, Kazan State Medical Academy, Kazan, Russia, 3 Research Laboratory "Omics technology", Institute of Fundamental Medicine and Biology, Kazan Federal University, Kazan, Russia, 4 Department of Microbiology and Immunology, University of Nevada, Reno, NV, United States, 5 Department of Pathology, University of Nevada, Reno, NV, United States*

Over 1,000 cases of hemorrhagic fever with renal syndrome (HFRS) were recorded in the Republic of Tatarstan (RT) in 2015. HFRS is a zoonotic disease caused by several different Old World hantaviruses. In RT, *Puumala orthohantavirus* (PUUV) is a prevalent etiological agent of HFRS. We looked for the genetic link between the PUUV strains isolated from the bank voles and from the infected humans. In addition, possible correlation between the genetic makeup of the PUUV strain involved and different clinical picture of HFRS was investigated. Partial PUUV small (S) genome segment sequences were retrieved from 37 small animals captured in the northwestern region of RT in 2015. Phylogenetic analysis revealed that 34 PUUV sequences clustered with strains of the previously identified "Russia" (RUS) genetic lineage, while 3 remaining PUUV sequences clustered with the known lineage from Finland (FIN). Sequence comparisons showed that the majority of the S-segment sequences isolated in the current study displayed 98.2–100.0% sequence identity when compared with the strains isolated earlier from the HFRS patients hospitalized in Kazan city. HFRS patients infected with PUUV strains of either RUS or FIN genetic lineages were observed to have consistent differences in clinical presentation of the disease and laboratory findings. These findings indicated a strong genetic link between the infected bank voles and human HFRS cases from the same localities. Thus, S-segment sequences of the PUUV strains isolated from HFRS patients could serve as a molecular marker for determining the likely geographic area where infection occurred.

Keywords: HFRS, RNA, PUUV, *Puumala orthohantavirus*, Republic of Tatarstan

#### *Edited by:*

*Hector A. Cabrera-Fuentes, University of Giessen, Germany*

#### *Reviewed by:*

*Sudheer Kumar Ravuri, Steadman Philippon Research Institute, United States Jan Clement, KU Leuven, Belgium Alexei B. Shevelev, Russian Academy of Sciences, Russia*

#### *\*Correspondence:*

*Sergey P. Morzunov<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Translational Pharmacology, a section of the journal Frontiers in Pharmacology*

*Received: 11 January 2019 Accepted: 29 July 2019 Published: 05 September 2019*

#### *Citation:*

*Davidyuk YN, Kabwe E, Shakirova VG, Martynova EV, Ismagilova RK, Khaertynova IM, Khaiboullina SF, Rizvanov AA and Morzunov SP (2019) Characterization of the Puumala orthohantavirus Strains in the Northwestern Region of the Republic of Tatarstan in Relation to the Clinical Manifestations in Hemorrhagic Fever With Renal Syndrome Patients. Front. Pharmacol. 10:970. doi: 10.3389/fphar.2019.00970*

## INTRODUCTION

The Republic of Tatarstan (RT) is among the regions of the Russian Federation (RF) with the highest annual load of the human cases of hemorrhagic fever with renal syndrome (HFRS). In 2015 alone, more than 1,000 cases of HFRS were diagnosed in the RT (Matrosov, 2016), thus placing this region into the fourth place with respect to the prevalence of this zoonosis infectious disease after Udmurtia, Bashkiria, and Mordovia in the Volga Federal District (VFD) of the Russian Federation (RF). *Puumala orthohantavirus* (PUUV) is one of the known causative agents of HFRS in humans, which causes a mild form of the disease with the fatality rate <0.43% (Lee, 1999; Khismatullina et al., 2016). PUUV belongs to the genus *Orthohantavirus*, family *Hantaviridae*. The main primary reservoir for PUUV is the bank vole, *Myodes glareolus* (Vapalahti et al., 2003; Tkachenko et al., 2012).

The PUUV tripartite genome consists of the S (small), M (medium), and L (large) single-stranded negative-polarity RNA segments: the S-segment codes for the nucleocapsid protein (NP), the most abundant protein produced in infected cells; the M-segment codes for the precursor of the envelope glycoproteins (Gn, Gc); and the L-segment codes for the RNA-dependent RNA polymerase (Plyusnin et al., 1996). Currently, eight genetic lineages of PUUV have been documented in various regions in Europe and western Siberia (Razzauti et al., 2013). Earlier, two distinct genetic lineages of PUUV were identified circulating within the bank vole populations in Russia. "Russia" (RUS) genetic lineage includes strains from the Samara region, Bashkiria, Udmurtia, and Tatarstan (Plyusnin et al., 1994; Lundkvist et al., 1997; Kariwa et al., 2009), and "Finland" (FIN) genetic lineage includes strains from Karelia and western Siberia (Asikainen et al., 2000; Dekonenko et al., 2003; Yashina et al., 2015). Between these two lineages, S-segment nucleotide sequences diversity reaches over 15% (Razzauti et al., 2012), while within a local rodent population, nucleotide sequence diversity does not exceed several percent within one lineage (Avsic-Zupanc et al., 2007; Kariwa et al., 2009; Razzauti et al., 2009). Although significant achievements have been made in the study of PUUV genetic diversity in Eurasia, only few investigations touched a question of the possible link between genetic composition of the PUUV strains in the bank voles and in the HFRS patients with different clinical manifestations of the disease (Horling et al., 1995; Plyusnin et al., 1997; Bahr et al., 2006; Garanina et al., 2009).

In our previous research (Davidyuk et al., 2017), we identified the PUUV strains in 25 patients diagnosed with HFRS in RT and 8 patients from the Republic of Mordovia. In the current study, genetic link between the PUUV strains isolated from the lungs of the bank voles and the RT PUUV strains identified earlier from infected humans was investigated. In addition, possible correlation was evaluated between genetic compositions of the PUUV strains infecting humans and differences in the clinical picture seen in the corresponding HFRS patients hospitalized in Kazan City.

### MATERIALS AND METHODS

*Patients.* Clinical manifestations of HFRS were studied in 74 patients who met the case definition and were receiving treatment in the Tatarstan Republican Clinical Infectious Diseases Hospital in Kazan City in 2015. According to the reports obtained from the patients, the infections occurred in the northwestern part of RT. Patients were hospitalized for 5–6 days of illness. The first sampling was carried out on the first day of admission to the clinic. Initial HFRS diagnosis based on the clinical and epidemiological criteria was confirmed by ELISA detection of antihantavirus IgM and IgG antibodies. The PUUV partial S-segment sequences were obtained from 25 patients in our previous work (Davidyuk et al., 2017). Three patients were excluded from the analysis because their clinical data were incomplete. The Institutional Review Board of the Kazan Federal University approved this study, and informed consent was obtained from each study subject according to the guidelines approved under the corresponding protocol.

*Tissue samples.* Frozen rodent lung tissue samples and information about trapping localities were obtained from Federal Healthcare Institution "Center for Hygiene and Epidemiology in the Republic of Tatarstan (Tatarstan)." Rodents were trapped in the northwestern regions of the Republic of Tatarstan in May and September 2015 (information about the geographic locations of the trapping sites is shown in **Figure 1** and **Table 1**). All PUUV sequences obtained from the serologically positive bank voles were named to include their corresponding virus name, trapping region, strain designation, and year (for example PUUV/ Vysokogorsky/MG\_054/2015).

#### RNA Extraction, cDNA Synthesis and PCR

Total RNA was extracted from lung tissues of bank voles with TRIzol Reagent kit (Invitrogen Life Technologies™, USA) following the manufacturer's recommendations. The concentration of RNA was determined with a NanoDrop 2000 UV–Vis spectrophotometer ("Thermo Fisher Scientific," USA). cDNA was synthesized using Thermo Scientific RevertAid Reverse Transcriptase ("Thermo Fisher Scientific," USA). PCR was performed using TaqPol polymerase kit ("Sileks," Russia) with primers PuuV-For 5-CTGCAAGCCAGGCAACAAACAGTGTCAGCA-3' and PuuV-Rev 5'-TCTGCCACATGATTTTTGTCAAGCACATC-3' (Kariwa et al., 2009). The resulting PCR products were purified with Isolate II PCR and Gel Kit ("Bioline," UK) and subsequently sequenced using ABI PRISM 310 big Dye Terminator 3.1 sequencing kit (ABI, USA). Obtained sequences were deposited in the GenBank database under accession nos. MG573266-573302.

### Phylogenetic Analysis

The nucleotide alignments and phylogenetic analysis of the PUUV strains based on the partial S-segment sequences 171 nucleotides in length (nucleotides 424–594) were performed with MegAlign program (Clustal W algorithm) located in the DNASTAR software package Lasergene (DNASTAR, USA) and MEGA v6.0 (Tamura et al., 2013). The parameters were adjusted manually.

PES, LAI-1, and LAI-2 (numbered 1–5, respectively).

TABLE 1 | Bank vole trapping site locations, number of trapped bank voles and nucleotide sequences obtained per location.

Phylogenetic trees were constructed using maximum parsimony method included in MEGA v6.0. For comparison, several partial S-segment sequences of the genetically distinct PUUV strains were downloaded from GenBank database (NCBI). These were included the following strains: Samara\_49/CG/2005, AB433843; Puu/Kazan, Z84204; PUUV Udmurtia/894Cg/91, Z21497; CG17/ Bashkiria-2001, AF442613; Sotkamo 2009, HE801633; PUUV/ Konnevesi/Mg\_O22B/2005, JQ319168; Kuchuk170/Mg/2007, KJ292966; and CRF366, AF367071. Tula virus strain Sennickerode Sen05/205, EU439951 was used as an outgroup.

#### RESULTS AND DISCUSSION

#### Identification and Sequence Comparisons of the PUUV Strains Found in the Bank Vole Populations in RT

Altogether, 129 bank voles were captured at 13 sites in May and September 2015. PUUV RNA was detected in 37 bank voles trapped at 11 sites, while viral RNA was not detected in 21 bank voles trapped at the sites S8 and S10 (**Table 1** and **Figure 1**). The proportion of infected bank voles in different sites vary. The high prevalence of infected bank voles was in sites S1–S4, S13, and S5–S7 located in two forests near the bank of the Volga river. In contrast, the lower proportion of infected animals was found in the isolated bank vole populations located in sites S9–S12 away from the Volga river. Plausible explanation is that bank voles are migration from sites S1–S4 to sites S5–S7. In addition, spatial distribution of hantavirus positive animals in different locations could be related to population densities and rates of virus transmission (Abbott et al., 1999; Mills et al., 1999). PUUV partial S sequences (564 base pairs, nt 242-805) were obtained from all reverse transcription PCR positive bank voles, analyzed and used for constructing phylogenetic trees. In addition, sequences obtained earlier from the strains circulating in the patients diagnosed with HFRS in 2015 (Davidyuk et al., 2017) and sequences of the selected PUUV strains from GenBank database were included in the analysis. The region of the S segment was selected based on the fact that it is shown to be one of the most variable region and often used for the analysis of genetic variability of hantaviruses (Asikainen et al., 2000; Sironen et al., 2001; Johansson et al., 2004; Klempa et al., 2006; Garanina et al., 2009; Ali et al., 2015). Sequence comparisons showed that PUUV sequences from each of the sites S1, S2, S4, S7, and S11– S13 displayed from 99.4 to 100.0% within-site nucleotide identity, although the sequences obtained from the site S3 were more variable (from 96.5 to 100% nucleotide identity). The sequences PUUV/Vysokogorsky/MG\_054/2015 and PUUV/Vysokogorsky/ MG\_058/2015 from the site S5 had 99.4% nucleotide identity while diverging significantly (83.6–84.8% nucleotide identity) from the rest of the sequences within the site S5 (see **Tables 1** and **2**). Nucleotide sequence identity between the sequences PUUV/Vysokogorsky/MG\_064/2015 and PUUV/Vysokogorsky/ MG\_066/2015 from the site S6 was as low as 83.0%.

Between-site nucleotide sequence comparisons revealed that 12 samples from the sites S1–S4 (sample pool from these four sites was designated as ZEL) shared 99.4–100.0% sequence identity. The only sequence that diverged slightly was PUUV/Zelenodolsky/ MG\_113/2015 (96.5–97.1% nucleotide identity with the rest of the ZEL pool). Likewise, the nucleotide sequences of 12 samples from the sites S5–S8 taken together (designed VYS-1 pool) displayed 98.8–100.0% nucleotide identity, and the nucleotide identity between the samples PUUV/Vysokogorsky/MG\_054/2015 and PUUV/Vysokogorsky/MG\_058/2015 from S5 and PUUV/ Vysokogorsky/MG\_064/2015 of S6 (designated VYS-2 pool) was 100.0%. The lowest nucleotide identity was observed between the sequences from VYS-1 and VYS-2 pools (83.0–84.2%).

The nucleotide sequences of the samples from the sites S11 and S12 (designated LAI-1 pool) were 100% identical. These samples also showed 95.9% nucleotide identity with the PUUV strains isolated from the site S13 (designated LAI-2 pool). The results of the comparison of the PUUV sequences from the different pools are shown in **Table 2**.

As expected, genetic variability of the PUUV S-segment nucleotide sequences obtained in the current study showed good correlation with geographic locations of the bank vole trapping sites. Sites S1–S4 are located in the western part of the sampled region, while sites S5–S8 are situated in the eastern part of the large forest (from Zelenodolsk to Vysokaya Gora), which borders the west and north of Kazan. There are no natural barriers to prevent migration of bank voles within the boundaries of this forest. Therefore, determining the boundaries of individual bank vole populations is difficult. All bank voles in this forest can be considered as one large population, which may include multiple local populations.

Likewise, sites S11 and S12 are located in the forest around Laishevo, and bank voles in this forest most likely form a distinct population; this conclusion is supported by the lower similarity observed between the PUUV sequences in the LAI-1 pool and in the ZEL/VYS pools. On the other hand, bank voles trapped in a relatively small isolated forest to the south of Pestretsy (S9) belong to a different population, and the PUUV strain identified in this forest differed significantly from the other groups.

The site S13 (LAI-2 pool) is located in the forest along the left bank of the Volga river west of Teteevo. The small mammals trapped there represent a group which is isolated from other populations included in the current study. As expected, corresponding PUUV sequences from LAI-2 pool are relatively distinct from ZEL/VYS pools and are most similar to those from LAI-1 pool.

Geographical locations of the trapping sites S9, S11, S12, and S13 and comparisons of the nucleotide identity values of the PUUV sequences from the corresponding PES, LAI-1, LAI-2 pools (**Figure 1** and **Table 2**) suggest that these three pools might be phylogenetically closer to each other than to ZEL and VYS pools.

Nucleotide sequence comparisons of the partial S-segment PUUV sequences with the corresponding sequences of the previously known PUUV strains found in GenBank showed the sequences from VYS-2 pool to be 100.0% identical to the corresponding nucleotide sequence of the PUUV strain Sotkamo and 93.0% identical to the sequence of the PUUV strain Konnevesi, both of the FIN lineage. The sequences of this pool displayed the lowest level of nucleotide identity (81.9–84.8%) with the strains of the RUS lineage previously known from the Volga region of Russia, such as Samara, Kazan, Udmurtia, and Bashkiria strains (**Table 3**). On the other hand, nucleotide sequences of ZEL, VIS-1, PES, LAI-1, and LAI-2 pools were more closely related to the RUS genetic lineage (91.2–97.7%

nucleotide identity) and more genetically distant from the sequences of the FIN genetic lineage (81.9–83.6% nucleotide identity) (**Table 3**). Hence, most of the PUUV strains identified in the current study belong to the RUS genetic lineage, while only three PUUV strains, PUUV/Vysokogorsky/MG\_054/2015, PUUV/Vysokogorsky/MG\_058/2015, and PUUV/Vysokogorsky/ MG\_064/2015, belong to the FIN genetic lineage. Thus, two PUUV genetic lineages were detected circulating in the bank vole populations in the forests north of Kazan between Yash-Ketch and Vysokaya Gora. The possibility of such co-circulation of two distinct PUUV genetic lineages in one territory was demonstrated earlier (Razzauti et al., 2009). Since the PUUV strains of the FIN lineage were found in the several widely distributed geographic locations including central and southern parts of Finland (Vapalahti et al., 1992; Plyusnina et al., 2012; Razzauti et al., 2013), Karelia (Asikainen et al., 2000), and western Siberia (Dekonenko et al., 2003), it could be suggested that these strains are distributed continuously from northern Europe (Finland) to at least western Siberia (Omsk region). In this case, forests in the Republic of Tatarstan, the Mari El Republic, and the Republic of Udmurtia could represent another zone of contact between the bank vole populations carrying PUUV strains of these two distinct genetic lineages. Consequently, the emergence of the reassortant and recombinant PUUV strains could happen in these areas. The verification of this assumption requires further investigation.

#### Phylogenetic Analysis of the Rodents and HFRS Patients' PUUV Strains

It is noteworthy that PUUV strains from PES, LAI-1, and LAI-2 pools showed slightly higher sequence identities with the corresponding sequences of Kazan and Udmurtia strains than the strains from ZEL and VYS-1 pools (see **Table 3**). Perhaps, this finding reflects the bank vole migration from the south to the north along the Volga River in the postglacial period (Dekonenko et al., 2003). Interestingly, only the PUUV sequences PUUV/ Vysokogorsky/MG\_054/2015, PUUV/Vysokogorsky/MG\_058/ 2015, and PUUV/Vysokogorsky/MG\_064/2015 demonstrated high identities (99.4–100.0%) to the strains, which were obtained earlier from HFRS patients infected with the strains belonging to FIN genetic lineage (Davidyuk et al., 2017). Nucleotide sequence identities between the PUUV strains of the RUS genetic lineage obtained from bank voles and the PUUV strains of the same lineage found in HFRS patients exceeded 90.0%. In particular, the comparisons revealed that most sequences of the PUUV strains from HFRS patients and the strains from bank vole pools (specifically, RT024, RT031, RT033, RT036, RT038, RT039, RT057 versus ZEL; RT048, RT058 versus LAI-1; RT055 versus LAI-2; RT002, RT005, RT006, RT008, RT010, RT011, RT013, RT014, RT043, RT050 versus VYS-2; and RT065 versus PES) share from 98.2 to 100.0% nucleotide identity. Based on these data, one could assume with high confidence that corresponding patients acquired hantavirus infection in the northwestern region of the Republic of Tatarstan and even determine the approximate location where infection occurred. In other words, the sequences of the PUUV strains isolated from the HFRS patients could be used as a molecular marker for determining the probable area of infection (**Figure 1**). Recently, similar data were published using HFRS cases diagnosed in some Europian countries, such as Germany, Finland, and Belgium (Plyusnin et al., 1997; Escutenaire et al., 2001; Ettinger et al., 2012). The data obtained may be used in the future for the development of the epidemiological measures aimed at preventing infection and reducing the incidence of HFRS in RT.

Surprisingly, one HFRS patient-originated PUUV strain RT012 did not show close genetic relationship to the strains identified in bank voles. This sequence displayed from 90.1 to 93.0% nucleotide identities with the sequences of the RUS genetic lineage isolated from bank voles and was 98.8% identical with "Bashkiria" strain. The patient might have contracted HFRS in the Volga region within the area where "Bashkiria" strain circulates including possibly unexplored regions of RT.

Phylogenetic tree of the PUUV strains obtained in the current study was reconstructed on the basis of the partial S-segment sequences (171 bp, nt 424-594) isolated from bank voles and HFRS patients (**Figure 2**). The PUUV nucleotide sequences obtained formed two distinct clades, which corresponded to the FIN and RUS genetic lineages. The RUS clade included the LAI-1, LAI-2, PES, and ZEL+VYS-1 subclades, containing isolates from RT and Bashkiria. Each subclade includes PUUV strains isolated from bank voles and HFRS patients. It should be noted that strains of the "ZEL+VYS-1" subclade are equidistant to all previously known RUS lineage.

The majority of the PUUV sequences obtained from bank voles (34 from 37) clustered with the RUS lineage, while the remaining three sequences from VYS-2 pool clustered with the FIN lineage. Interestingly, sequences obtained from the HFRS patients showed different pattern: 12 sequences clustered with the RUS lineage, while 10 sequences were placed in the FIN lineage. Relatively high proportion of the patients infected with

TABLE 3 | Nucleotide sequence identity of the FIN and RUS genetic lineages to the PUUV strains discovered in the northwest region of RT.

The compressed branches marked "hum + ZEL," "VYS-1," and "hum + VYS-2" includes the following: (i) strains RT024, RT031, RT033, RT036, RT038, RT039, and RT057 and nine strains from the bank voles trapped in sites S1–S4; (ii) 12 strains from VYS-1 pool; (iii) strains RT002, RT005, RT006, RT008, RT010, RT011,

the FIN lineage strains could possibly be explained by: i) such strains circulate in the bank vole populations in the northwestern territory of RT that have not been sampled yet; ii) the infections could have occured within the limited geographic area around Yash-Ketch and Vysokaya Gora. More genetic analysis of PUUV circulating in RT and neighboring regions is required to explain variations in the number of patients infected with FIN and RUS virus lineages.

RT013, and RT 014 and three strains from VYS-2 pool, and strain Sotkamo 2009, respectively.

### Different Clinical Manifestations in the Patients With HFRS Caused by the PUUV Strains of the FIN and the RUS Lineages

In order to compare clinical manifestations observed in the HFRS patients infected with the PUUV strains of the RUS versus FIN genetic lineages, clinical and laboratory data from 22 patients (17 male and 5 female) were analyzed. The patients were divided in two groups: group 1 included 12 patients from whom the PUUV strains of the RUS lineage were recovered (group "RUS"), and group 2 consisted of 10 patients infected with the PUUV strains of the FIN lineage (group "FIN"). The average age of the patients was 35.9 ± 12.06 years in "RUS" group and 49.8 ± 14.6 years in "FIN" group, respectively (*p* > 0.05). The average hospitalization period and the febrile and oliguric periods did not differ significantly in both groups (**Table 4**). Urinalysis revealed proteinuria in 10 and 6 cases (*р* > 0.05) for "RUS" and "FIN" groups, respectively. Hematuria was observed in six "RUS" group and four "FIN" group cases (*р* > 0.05). Bleeding in the form of scleral hemorrhage was revealed in only one patient from "RUS" group. The kidney edema was observed in nine patients of the "RUS" group and in five patients TABLE 4 | Clinical and laboratory characteristics of HFRS patients, infected with PUUV strains of "RUS" versus "FIN" genetic lineages.

*aUltrasound diagnosis.*

*bOliguria— < 500 ml of urine for 1–3 days.*

*cProteinuria— > 30 mg/dl of protein in urine.*

*dHematuria— > 60 red blood cells per high-power microscopic field.*

*eThe bolded p values are statistically significant to clinical manifestations in different groups.*

of the "FIN" group (*р* > 0.05). Increased serum levels of urea were detected in 5 out of 12 in "RUS" and 3 out of 10 cases in "FIN" groups (*р* > 0.05).

Furthermore, five of the patients from "RUS" group and four from "FIN" group experienced acute lung injury (*р* > 0.05). Gastrointestinal (GA) symptoms including stomach pain, nausea, vomiting, and diarrhea were documented in seven "RUS" group and three in "FIN" group (*р* > 0.05), while oliguria was found in nine "RUS" group and three "FIN" group of HFRS cases (*р* > 0.05). Impaired vision was observed only in the "RUS" PUUV lineage infected patients (4, *p* < 0.05), while pain in the lumbar region was experienced by 11 of the "RUS" patients and by 5 of the "FIN" patients (*p* < 0.05) (**Table 4**). Among the laboratory parameters investigated, patients of the "RUS" group showed an increased level of urea and creatinine as compared to the "FIN" group: 12.92 ± 3.20 mmol/l vs. 6.25 ± 0.82 mmol/l (*р* > 0.05) and 208.08 ± 56.14 μmol/l vs. 104.90 ± 3.52 μmol/l (*p* > 0.05) (**Table 4**). The "RUS" PUUV lineage-infected patients had a significantly lower platelet count (72.16 ± 12.16 × 109 /l) and higher aspartate aminotransferase (AST) level (50.83 ± 7.77 U/l) as compared to the "FIN" group, where thrombocyte counts were 131.0 ± 14.71 × 109 /l (*p* < 0.05) and AST level was 31.6 ± 3.76 U/l (*p* < 0.05).

Overall, the clinical manifestations seen in the HFRS patients of both groups mentioned above matched typical HFRS clinical manifestation caused by PUUV and were in agreement with the description given in the works of many authors (Trusov et al., 2004; Shakirova et al., 2011; Rasmuson et al., 2013). HFRS was characterized by acute onset with fever, malaise, headache, pain in the eyes, and abdominal pain. Acute kidney injury diagnosed in 40–100% of patients depending on the disease severity. Hemorrhages are detected in some HFRS cases, including petechia, gastrointestinal, and nasal bleeding. Acute lung injury is evident in some patients and often characterized by mild to moderate dyspnea or dry cough (Rasmuson et al., 2013). In severe cases, lung edema was found (Trusov et al., 2004; Rasmuson et al., 2013). However, the significant differences in the course of the disease and biochemical findings observed in the current study suggest that the PUUV strains of the RUS lineage cause a disease with more severe clinical symptoms, including more pronounced hemorrhages and renal manifestations, unlike the PUUV strains of the FIN lineage, which cause a milder form of HFRS.

We believe that the results of this investigation are of particular importance not only for the local health authorities in Tatarstan but also for the international scientific and public health community. As PUUV could be found throughout most of Europe and Asia, PUUV strains currently circulating in RT are integral part of the Old World HFRS epidemics. Our current data confirm that PUUV strains circulating in RT (called Finnish and Russian genetic lineages) are genetically related to the virus strains found in the Western and Central Europe. Thus, current data improve our understanding of PUUV distribution and genetic diversity. Ongoing climate change could influence the bank vole migration patterns and therefore affect future distribution and genetic composition of PUUV strains throughout its entire geographic distribution. PUUV strains could "migrate" together with their natural hosts into the new areas that were not endemic in the past. In addition, increased international trade could inadvertently promote introduction of PUUV rodent hosts into the new areas. Thus, comprehensive understanding of PUUV epidemiology in RT could assist in predicting the virus spread beyond its current geographic range and investigating future HFRS outbreaks.

### ETHICS STATEMENT

The Ethics Committee of the Kazan Federal University approved this study and informed written consent was obtained from each NE patient and controls according to the Guidelines approved under this Protocol (article 20, Federal Law "Protection of Health Right of Citizens of Russian Federation" N323- FZ, 11.21.2011).

## AUTHOR CONTRIBUTIONS

YD and SK performed experiments; EM, EK, and VS made the conceptualization and data curation; AR and IK analyzed data; SM, and RI analyzed data and wrote the manuscript.

### ACKNOWLEDGMENTS

The work was performed as a part of the Russian Government Program of Competitive Growth for the Kazan Federal University. Some of the experiments were conducted using the equipment of the Interdisciplinary Center of Shared Facilities and the Scientific and Educational Center of Pharmaceutics of Kazan (Volga Region) Federal University, Kazan, Russian Federation. Albert A. Rizvanov was supported by state assignments 20.5175.2017/6.7 and 17.9783.2017/8.9 of the Ministry of Science and Higher Education of Russian Federation.

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2019 Davidyuk, Kabwe, Shakirova, Martynova, Ismagilova, Khaertynova, Khaiboullina, Rizvanov and Morzunov. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Surveillance of Tumour Development: The Relationship Between Tumour-Associated RNAs and Ribonucleases

*Nadezhda Mironova\* and Valentin Vlassov*

*Institute of Chemical Biology and Fundamental Medicine, Siberian Branch of Russian Academy of Sciences, Novosibirsk, Russia*

Tumour progression is accompanied by rapid cell proliferation, loss of differentiation, the reprogramming of energy metabolism, loss of adhesion, escape of immune surveillance, induction of angiogenesis, and metastasis. Both coding and regulatory RNAs expressed by tumour cells and circulating in the blood are involved in all stages of tumour progression. Among the important tumour-associated RNAs are intracellular coding RNAs that determine the routes of metabolic pathways, cell cycle control, angiogenesis, adhesion, apoptosis and pathways responsible for transformation, and intracellular and extracellular non-coding RNAs involved in regulation of the expression of their proto-oncogenic and oncosuppressing mRNAs. Considering the diversity/variability of biological functions of RNAs, it becomes evident that extracellular RNAs represent important regulators of cell-to-cell communication and intracellular cascades that maintain cell proliferation and differentiation. In connection with the elucidation of such an important role for RNA, a surge in interest in RNA-degrading enzymes has increased. Natural ribonucleases (RNases) participate in various cellular processes including miRNA biogenesis, RNA decay and degradation that has determined their principal role in the sustention of RNA homeostasis in cells. Findings were obtained on the contribution of some endogenous ribonucleases in the maintenance of normal cell RNA homeostasis, which thus prevents cell transformation. These findings directed attention to exogenous ribonucleases as tools to compensate for the malfunction of endogenous ones. Recently a number of proteins with ribonuclease activity were discovered whose intracellular function remains unknown. Thus, the comprehensive investigation of physiological roles of RNases is still required. In this review we focused on the control mechanisms of cell transformation by endogenous ribonucleases, and the possibility of replacing malfunctioning enzymes with exogenous ones.

Keywords: tumour-associated RNA, extracellular miRNAs, tumour development, ribonucleases, RNA degradation

## TUMOUR-ASSOCIATED RNAS AND THEIR ROLE IN CARCINOGENESIS

Tumour development is accompanied by rapid cell proliferation, loss of differentiation, the reprogramming of energy metabolism, loss of adhesion between tumour cells and matrix, evasion of immune surveillance, angiogenesis induction, infiltration growth, and metastatic spreading (Hanahan and Weinberg, 2011). Tumour-associated RNAs play an important role at all stages

#### *Edited by:*

*Hector A. Cabrera-Fuentes, University of Giessen, Germany*

#### *Reviewed by:*

*Karla Mayolo-Deloisa, Monterrey Institute of Technology and Higher Education (ITESM), Mexico Ester Boix, Autonomous University of Barcelona, Spain*

> *\*Correspondence: Nadezhda Mironova<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Translational Pharmacology, a section of the journal Frontiers in Pharmacology*

*Received: 19 April 2019 Accepted: 09 August 2019 Published: 13 September 2019*

#### *Citation:*

*Mironova N and Vlassov V (2019) Surveillance of Tumour Development: The Relationship Between Tumour-Associated RNAs and Ribonucleases. Front.  Intracellular and extracellular non-coding RNAs participate in the regulation of the expression of target proto-oncogenic and oncosuppressive mRNAs as well (Dalmay and Edwards, 2006; Esquela-Kerscher and Slack, 2006).

Different groups unequivocally demonstrated the presence of various RNAs, including tumour-derived and tumour-associated RNAs, in the blood samples of patients with oncological diseases (Wieczorek et al., 1985; Kopreski et al., 1999; Vlassov et al., 2010). The range of RNAs discovered in blood plasma samples is rather wide and includes fragments of ribosomal RNA (rRNA), messenger RNA (mRNA), transport RNA (tRNA), mitochondrial RNA, small non-coding RNA particularly miRNA being detected in plasma or serum in RNA/protein complexes and within extracellular membrane vesicles (EV) (Valadi et al., 2007; Savelyeva et al., 2016; Savelyeva et al., 2017).

Data are accumulated that EV of cancer origin comprise pathogenic components, including mRNA and miRNA, that together with other components such as DNA proteins, transcriptional factors and lipids can take part in paracrine signalling in the tumour microenvironment (Fujita et al., 2016). These evidences also predict the role of EV-mediated transfer of cancer-associated biomolecules to distant organs contributing to the initiation of pre‐metastatic niche formation. In support of this, an increasing number of publications has demonstrated the contribution of EV RNA in events of cancer development, namely, cell proliferation (Hong et al., 2009; Kogure et al., 2011; Zhang et al., 2015a), drug resistance (Challagundla et al., 2015), angiogenesis (Kosaka et al., 2013), immune modulation (Fabbri et al., 2012), and pre‐metastatic niche formation (Fong et al., 2015).

### Extracellular Circulating mRNAS

Among the extracellular miRNA (ex-miRNA), mRNA fragments encoding tumour-associated antigens were detected, i.e. proteins that are expressed at low level in the cell, but overexpressed upon tumour progression.  Overexpression of telomerase in cells can lead to epithelial-mesenchymal degeneration and tumour progression.

High levels of mRNA encoding mammaglobin, CK-19 (keratin 19) (Silva et al., 2001) and HER2/neu (Erb-B2 receptor tyrosine kinase 2) (Nicolini et al., 2017), which are specific markers of breast cancer (Lianidou et al., 2015), are also found in the blood plasma of patients with given disease. CK-19 is involved in maintaining the stability of epithelial cells and high expression levels are detected in tumour tissue (Lianidou et al., 2015). The HER2/neu oncogene is amplified at a high level in approximately 20% of all breast cancer cases and is related to rapid tumour proliferation and a poor prognosis.

In the case of lung cancer, the elevated levels of mRNA fragments encoding HER2/neu, hnRNP-B1 (heterogeneous nuclear ribonucleoproteins B1) and 5Т4 (oncofetal trophoblast glycoprotein) were detected in blood (Kopreski et al., 2001; Fleischhacker et al., 2001; Sueoka et al., 2005). hnRNP-B1 was found to play a significant role in the splicing of tumour suppressors and is considered an oncogene. It has also found to be crucial in the development of glioblastoma, hepatocellular carcinoma and lung cancer (Zech et al., 2006; Golan-Gerstl et al., 2011; Shilo et al., 2014). Oncogene 5T4 is involved in the modulation of cell adhesion and is over-expressed in many types of cancer cells, as along with such oncogenes as MYC (protooncogene c-Myc) and NRAS (NRAS proto-oncogene, GTPase).

For patients with malignant melanoma, the presence in blood of mRNA encoding for tyrosinase, MAGE-3 (melanomaassociated antigen 3), MCAM (melanoma cell adhesion molecule), p97 (transitional endoplasmic reticulum ATPase) and HMBS (hydroxymethylbilane synthase) is typical (Hoon et al., 2000; Hasselmann et al., 2001). Tyrosinase is involved in the synthesis of melanin, MAGE-3 is involved in malignant transformation and is the main tumour-specific melanoma antigen, and MCAM is involved in cell adhesion.

## Circulating Non-Coding RNAs

The pool of non-coding RNAs discovered in the bloodstream consists of long non-coding RNAs (lncRNA), short noncoding RNA including microRNAs (miRNA), and the recently discovered piwi-interacting RNAs. Long non-coding RNAs and piwi-interacting RNAs are described in detail in a recent review (Rapisuwon et al., 2016). Here we will focus on ex-miRNAs, which are important in the regulation of tumour development.

 In blood plasma, the main component of the ex-miRNAs are preserved enclosed in the ribonucleoprotein complexes comprising Ago2 protein, microvesicles and apoptotic bodies, and high and low-density lipoproteins (Valadi et al., 2007; Hunter et al., 2008; Zernecke et al., 2009; Vickers et al., 2011; Turchinovich et al., 2011; Arroyo et al., 2011; Bayraktar et al., 2017; Pucci et al., 2018). In this regard, miRNAs are highly stable and resistant to blood ribonucleases and can be transferred throughout the body, which makes their regulatory function virtually unlimited.

It is worth mentioning that miRNAs regulate fundamental cellular processes, such as proliferation, differentiation, metabolism, DNA repair, apoptosis, and transformation and can function as mediators in cell-to-cell communication, thereby acting like hormones (Ambros, 2004; Bartel, 2004; Croce and Calin, 2005; Calin and Croce, 2006; Cortez et al., 2011; Anfossi et al., 2018).

Circulating ex-miRNAs include both oncomirs and oncosuppressors.   Other miRNAs can also be indicative of other forms of cancer: miR-125a-3p – colon cancer (Wang et al., 2017), miR-200c and miR-141 – metastatic breast cancer (Zhang et al., 2017a); and the miR-200 family – prostate cancer and high-grade serous epithelial ovarian cancer (Lin et al., 2017; Pendlebury et al., 2017).

miR-21 represents one of the first microRNAs being defined as an oncomir, that regulate multiple tumour suppressors like the PTEN (phosphatase and tensin homolog), PDCD (programmed cell death), p53 (tumor suppressor p53) and TP63 (tumor protein p63) pathways (Meng et al., 2007; Asangani et al., 2008; Papagiannakopoulos et al., 2008). miR-155 act as an oncogene by inhibition of suppressor of cytokine signaling 1 (SOCS1) expression (Jiang et al., 2012). Taking into account the wellknown functional link between inflammation and cancer, and the fact that inflammation to some extent is mediated by miR-155, the oncogenic role of miR-155 becomes clear (Jiang et al., 2010). However, although there is a lot of evidence for the oncogenic role of miRNA-155, it can also act as a tumour suppressor (Higgs and Slack, 2013).

Besides oncomirs, increased levels of which are identified in the blood of patients with various oncological diseases, tumour suppressor miRNAs can also be found in the blood stream. In this regard two or more miRNAs are used as diagnostic (by the level of oncomirs) and prognostic (by the level of tumour suppressor miRNAs) markers (Yang et al., 2015; Elghoroury et al., 2018).

The levels of tumour suppressor miRNAs are usually decreased in the blood of cancer patients, such as miRNAs belonging to let-7 family. Specifically, let-7 miRNAs directly interact with mRNAs encoding proteins involved in the cell cycle and signal transduction pathways that lead to carcinogenesis (Büssing et al., 2008). The decreased expression of let-7b is usually observed in lymph node metastases of breast cancer cells (Thammaiah and Jayaram, 2016). In addition, down-regulation of let-7b/g is evidenced during gastric cancer development being associated with poor survival and lymph node metastasis (Kang et al., 2014). A decreased level of miR-152 has been also detected in various human cancer cell lines and tumour tissues, such as gastrointestinal (Chen et al., 2010), endometrial (Tsuruta et al., 2011) and ovarian cancer (Zhou et al., 2012), as well as hepatocellular carcinoma (Huang et al., 2010), indicating that miR-152 might act as a tumour suppressor in these tumours.

### Role of Tumour-Associated Extracellular RNA (ExRNA) in Transformation

The secretion of RNAs in ribonucleoprotein complexes (RNP) by cells, and the transfer of those RNP between mammalian cells, was for the first time established in the 1970s (Kolodny et al., 1972). RNAs in RNPs can be a specific product released from tumour cells that may mediate host-tumour interaction and regulation of gene expression (Rosenberg-Nicolson and Nicolson, 1994). The recent discovery of regulatory RNAs, particularly miRNAs, has led to a revolutionary hypothesis that ex-miRNAs can mediate cell-to-cell signalling by paracrine or even endocrine manner, especially playing a crucial role in the context of cancer and metabolism. This hypothesis arose because several research groups found a large amount of miRNA in the bloodstream and was largely supported by numerous subsequent publications demonstrating that miRNAs in RNPs or EV can enter the recipient cells, change gene expression, and cause functional effects (Valadi et al., 2007; Skog et al., 2008; Mittelbrunn et al., 2011; Montecalvo et al., 2012).

Cell-to-cell communication by ex-miRNA has also been proven for cells of the immune system. Exosomal-transfered pro-inflammatory miR-155 and immunosuppressive miR-146a from dendritic cells was demonstrated to reduce the level of their mRNA targets, and reprogrammed the response of cellsrecipients to endotoxin (Alexander et al., 2015). The mechanism of regulation of the activity and differentiation of mast cells was shown to be mediated by the transfer of mRNA and miRNA in exosomes between cells (Valadi et al., 2007). Recent studies have demonstrated that adipose-derived EV-circulating miRNAs participate in cell–cell crosstalk between adipose and liver tissues by altering mRNA expression and translation in target tissue (see review Thomou et al., 2017).

Communication between tumour and normal cells provides a route for tumours to manipulate their environment, making it more favourable for growth and invasion.  One example is the destruction of tight contacts in the epithelial cells of blood vessels under the action of exosome-derived miR-105 secreted by neighbouring cancer cells, and the subsequent increase in metastasis (Zhou et al., 2014). EV-derived miRNA-21 secreted by hepatocellular carcinoma cells participates in tumour progression triggering the conversion of hepatocyte stellate cells to cancerassociated fibroblasts (Zhou et al., 2018). MiR-210 obtained from hepatocellular carcinoma cells as well, was found to promote endothelial cell migration along with angiogenesis, which was confirmed by the correlation between the elevated level of miR-210 in the blood of patients with hepatocellular carcinoma and high microvessel density (Lin et al., 2018). Thus, tumour-derived miRNAs are the tools for the transformation of the normal cells to malignant, and adjustment of their microenvironment for favorable tumor development.

### THE ROLE OF ENDOGENOUS MAMMALIAN RNASES IN THE CONTROL OF INTRACELLULAR EVENTS AND SIGNALLING PATHWAYS RESPONSIBLE FOR TRANSFORMATION

Benner hypothesized that a certain balance between RNAs, RNases, and ribonuclease inhibitors controls tissue development in higher organisms (Benner, 1988). Many factors participated in regulation of gene expression at the mRNA level. These factors are non-coding RNAs, RNA-binding proteins and RNases that maintain RNA-homeostasis in cells and discard aberrant RNAs through the degradation and turnover of transcripts, control of RNA decay, and biogenesis of miRNAs. Disruption of mentioned processes is mainly associated with distortion of expression or the improper functioning of these factors followed by cell transformation and tumour development. Among these factors, RNases play quite important role since they regulate the turnover of various transcripts at every stage of the cell cycle and participate in the processing of RNA involved in translation control.

In a number of studies up to 2009, it was shown that intracellular RNases are involved in both induction and suppression of tumour progression (see review Kim and Lee, 2009). Nowadays a lot of information has appeared expanding the supervisory function of exogenous ribonucleases in the RNA world (**Tables 1** and **3**) and their intracellular RNA-targets (**Figure 1**).

#### RNases of Conventional RNA Decay

Deadenylation is an essential way of regulation of mRNA stability and expression of genes responsible for the fundamental functions such as development and differentiation at cell level under normal or pathological conditions, including chronic inflammation and cancer (Zhang et al., 2010; Zhang et al., 2015b). The CCR4-NOT complex, a major deadenylase in mammals, plays dual roles in the control of tumour development. The mammalian CCR4-NOT complex was described to comprise eight subunits: CNOT1, CNOT2, CNOT3, CNOT6 or 6L, CNOT7 or 8, CNOT9, CNOT10, and CNOT11 with four of them, namely, CNOT6/6L/7/8 and CNOT3 exhibited deadenylase properties (Bartlam and Yamamoto, 2010; **Figure 1B**).

Knockdown of CNOT3, a subunit incorporated in the CCR4- NOT complex and responsible for deadenylase activity, was shown to induce tumour development in a sensitized drosophila eye cancer model (Vicente et al., 2018). Moreover, mutations of the CNOT3 gene were discovered in samples of T-cell acute lymphoblastic leukemia (T-ALL) patients (De Keersmaecker et al., 2013), suggesting the tumour suppressor role of CNOT3. However, contrary to the tumour suppressive function components of the CCR4-NOT complex, it can have an impact on cancer progression. For instance, the up-regulation of CNOT3 promotes the progression of non-small cell lung cancer (Shirai et al., 2019) and activity of CNOT7 may stimulate migration of mouse breast cancer cells (Faraji et al., 2016; **Table 3**).

It is known that deadenylases facilitate miRNA-induced mRNA decay resulted from their interaction with the miRNA-induced silencing complex (miRISC). A vast amount of publications give an evidence of participation of deadenylation complexes, such as CCR4-NOT and Pan2–Pan3, in miRNA-mediated deadenylation being necessary for regulation of gene expression and stability of mRNA (see review Zhang et al., 2010). Poly(A)-specific ribonuclease (PARN) is an important deadenylase: among its targets are migration and adhesion factors, as well as mRNAs of proteins involved in p53, FAK (fokal adhesion kinase), and MAPK signaling (Lee et al., 2012; Devany et al., 2013; **Figure 1A**). PARN may also participate in miRNA-mediated deadenylation due to association with Ago2 in the RNA-induced silencing complex (RISC), and promote degradation of the oncogenic miR-21 followed by restoration of tumor suppressor activity of corresponding protein targets such as PTEN and p53 (Boele et al., 2014; Zhang et al., 2015a). PARN inhibition was shown to induce p53 accumulation and decrease cancer cell viability (Shukla et al., 2019).

5′–3′ exonuclease XRN1 is an enzyme being involved in conventional RNA decay (Long and McNally, 2003), which is also implicated in cancer as a tumour suppressor (**Table 3**). The decreased expression and/or complete depletion of XRN1 mRNA were found in primary samples of osteogenic sarcoma (Zhang et al., 2002). XRN1 realized additional control over epithelial to mesenchymal transition (EMT) on the level of ex-miRNA decay. It was found that XRN1 degrades ex-miRNA-223 derived from extracellular vesicles of polymorphonuclear leukocyte neutrophils after penetration into tumour cells, thus promoting transient epithelial-mesenchymal transition (Zangari et al., 2017; **Figure 1C**). Recently obtained data show that XRN1 negatively regulates autophagy in mammalian cells that thus reduces cell survival, which reinforces the evidence that this is a suppressor (Delorme-Axford et al., 2018). Contrary to this, 5′–3′ exonuclease XRN2 promotes EMT and metastasis through regulation of the processing of pre-miR-10a to mature miR-10a, and is a candidate inducer of spontaneous lung cancer (Zhang et al., 2017b; **Figure 1C**).

### Stress Signal Induced RNases

A number of endogenous RNases (RNase L, IRE1α, and PMR1) are normally silent in the cell and are induced under specific stress signals to effect tumour-modulating functions. Human RNase L displaying endoribonuclease activity expressed in many types of normal and cancerous mammalian cells (Zhou et al., 2005). RNase L is single-stranded ribonuclease able to cleave viral RNA, rRNA, and some cellular RNA both in cells and cell-free systems, at phosphodiester bonds in UU and UA sequences (Wreschner et al., 1981; Li et al., 1998; **Figure 1D**). In human and mouse cells, RNase L controls the stability of mRNA encoded in mitochondria and destabilizes the mRNA of genes induced by the interferon response to a viral infection (Li et al., 1998; Le Roy et al., 2001). RNase L is normally involved in innate immunity and antiviral defence

(Malathi et al., 2007), however besides these functions it also plays a role as a tumour suppressor. Mutations in the RNase L gene were found to contribute to enhanced cell migration and invasion, and knockdown of RNase L in human prostate cancer cell line PC3 resulted in increase of tumour growth rate and metastases spreading *in vivo* (Banerjee et al., 2015; Dayal et al., 2017; **Table 3**). Cleavage of mRNAs encoding proteins involved in cell adhesion and migration appears a more likely mechanism for the inhibition of cell migration by RNase L (Banerjee et al., 2015). Interestingly, RNase L can discriminate and eliminate exogenous miRNA mimics (Nogimori et al., 2019; **Figure 1D**).

IRE1α is a serine/threonine kinase, an endoribonuclease, which is one of the major participants in endoplastic reticulum (ER) proteostasis and plays a dual role in cancer development (**Table 3**). It carries out both tumour-inducing and tumoursuppressing activity. Activation of IRE1α was observed in several types of tumors and was associated with overexpression of such oncogenes as BRAFV600E (mutant form V600E of B-Raf proto-oncogene, serine/threonine kinase gene), MYC, and HRAS (HRAS proto-oncogene, GTPase) (Croft et al., 2014). In turn, activation of IRE1α and its functioning as ribonuclease may lead to the process named RNA regulated IRE1‐dependent decay (RIDD) that represent degradation of mRNA and miRNA targets (Maurel et al., 2014). In mammalian cells, the substrates for IRE1α are its own mRNA, mRNA encoding XBP1 and CD59, and other mRNAs encoding proteins involved in the regulation of angiogenesis (see review Kim and Lee, 2009; **Figure 1E**).

Several studies demonstrated that inhibition of the expression or the RNase activity of IRE1 suppresses the development of several types of tumours, mostly because of the ablation of prosurvival effects of XBP1 on tumour growth (Chevet et al., 2016; Obacz et al., 2017). Recently inhibition of IRE1 ribonuclease activity was found to influence the tumour cell secretome and enhance its sensitivity to chemotherapy (Logue et al., 2018). The tumour suppressive function of IRE1 was also detected. In several studies on genome screening, it was found that IRE1α is often found in the mutant form in various types of malignancies (Parsons et al., 2008; Guichard et al., 2012). Overexpression of IRE1 leads to a decrease in the expression of CD59, being implicated in the progression of lung cancer (Oikawa et al., 2007). Thus, IRE1 is an important RNase that exhibits a dual role in cancer progression by directing cancer progression and cell death.

PMR1 exhibits the properties of a proto-oncogene and is an effector of the EFGR (epidermal growth factor receptor) signalling pathway. Recently obtained data shows that increased migration activity and invasiveness of MCF-7 breast cancer cells is associated with high PMR1 activity, the targets of which are miRNAs of the miR-200 family, which are responsible for controlling adhesion and invasion (Bracken et al., 2014; Gu et al., 2016; Perdigão-Henriques et al., 2016; **Figure 1F**).

#### Proteins Regulating mRNA Stability

RAS-GTPase-activating protein (SH3 domain)-binding proteins (G3BPs) represent a family of proteins capable of RNA binding and able to regulate mRNA stability and translation in response to environmental stresses (**Table 1**). The mammalian G3BP family consists of homologous proteins G3BP1, G3BP2a, and its splice variant G3BP2b with a similar molecular structure, which are located in the nucleus and cytoplasm. The different functions of G3BPs are summarized in a range of reviews (see revs Kim and Lee, 2009; Alam and Kennedy, 2019). From the point of view of its influence on the RNA world, it is important to note that GB3P1 participates in RNA metabolism including regulation of various cellular mRNAs and miRNAs. G3BP1 controls certain transcripts either due to its ability to stabilize mRNA like mRNA *tau* and *CDK7* (cyclin dependent kinase 7) (Atlas et al., 2004) or to cause mRNA degradation as in the case of mRNA MYC, BART (Epstein-Barr virus derived RNA encoding a set of miRNAs), *CTNNB1* (catenin beta 1), *PMP22* (peripheral myelin protein 22), *GAS5* (growth arrest specific 5), and *IGF2* (insulin like growth factor 2) (Gallouzi et al., 1998; Tourrière et al., 2001; Zekri et al., 2005; Taniuchi et al., 2011a; Taniuchi et al., 2011b; Winslow et al., 2013; **Table 3**). In earlier papers G3BPs were suggested to play a role in tumour development since their elevated levels were found in different types of proliferating cells and tumours (Guitard et al., 2001; Barnes et al., 2002; French et al., 2002). Moreover, G3BPs were found to be participants of key cell-growth associated molecular pathways important for tumorigenesis including RAS, the NF-κB, and MAPK pathways, and the ubiquitin proteasome system (Gallouzi et al., 1998; Prigent et al., 2000; Soncini et al., 2001; **Table 3**).

#### Proteins With Ribonuclease Activity Participating in Maintenance of DNA Integrity

In addition, it was found that enzymes involved in DNA replication and repair, such as APE1, and FEN1, also exhibit RNase activity. Under stress, or when a nuclear localization

TABLE 1 | Endogenous RNases and other proteins with ribonuclease activity participating in maintenance of normal RNA homeostasis of eukaryotic cells.

*CNOT3, CCR4-NOT transcription complex subunit 3; CNOT7, CCR4-NOT transcription complex subunit 7; PARN, poly(A)-specific ribonuclease; XRN1, 5'-3' exoribonuclease 1; XRN2, 5'-3' exoribonuclease 2; RNase L, ribonuclease L; IRE1a, serine/threonine-protein kinase/endoribonuclease IRE1; PMR1, ATPase secretory pathway Ca2+ transporting 1; ANG, angiogenin, ribonuclease, RNase A family, 5; G3BP1, GTPase activating protein (SH3 domain) binding protein 1; APE1, apurinic/apyrimidinic endodeoxyribonuclease 1; FEN1, flap structure-specific endonuclease 1; Drosha, double-stranded RNA-specific endoribonuclease, nuclear; Dicer, double-stranded RNA-specific endoribonuclease, cytoplasmic; Ago2, Argonaute RISC catalytic component 2.*

signal is lost, these enzymes redistribute between the nucleus and the cytoplasm, where they can affect the level of cellular RNA. Two enzymes, APE1 and FEN1, have recently attracted close attention because of their ability to cleave RNA and the fact that their expression is associated with oncogenesis.

Apurinic/apyrimidinic endodeoxynuclease 1 (APE1) is an enzyme that exhibits both deoxyribonuclease and ribonuclease activity (**Table 1**). APE1 is mainly associated with DNA repair and redox regulation of transcription factors. In base excision repair (BER), APE1 functions as an apurinic/apyrimidinic endodeoxyribonuclease and corrects DNA damage caused by oxidizing or alkylating agents. APE1 was also found to exhibit endoribonuclease activity targeting *MYC* mRNA (Barnes et al., 2009; **Figure 1H**), and cleaving several other RNAs at UA, UG, and CA sites in the single stranded regions *in vitro* (Bergstrom et al., 2006). APE1 participates in rRNA quality control processes during cell division (Vascotto et al., 2009). Thus, APE1 performs several functions in the cell and can encourage genetic integrity and modulate turnover of different mRNAs as a ribonuclease. Recently, it has been suggested that this protein can perform non-canonical, but, nevertheless, important functions in RNA metabolism, regulating post-transcriptional expression of genes (Tell et al., 2010; Antoniali et al., 2014; Antoniali et al., 2017a).