*Article* **Oncolytic H-1 Parvovirus Hijacks Galectin-1 to Enter Cancer Cells**

**Tiago Ferreira 1, Amit Kulkarni 2, Clemens Bretscher 1, Petr V. Nazarov 3, Jubayer A. Hossain 4, Lars A. R. Ystaas 4, Hrvoje Miletic 4,5, Ralph Röth 6,7, Beate Niesler 6,7 and Antonio Marchini 1,2,\*,†**


**Abstract:** Clinical studies in glioblastoma and pancreatic carcinoma patients strongly support the further development of H-1 protoparvovirus (H-1PV)-based anticancer therapies. The identification of cellular factors involved in the H-1PV life cycle may provide the knowledge to improve H-1PV anticancer potential. Recently, we showed that sialylated laminins mediate H-1PV attachment at the cell membrane. In this study, we revealed that H-1PV also interacts at the cell surface with galectin-1 and uses this glycoprotein to enter cancer cells. Indeed, knockdown/out of *LGALS1,* the gene encoding galectin-1, strongly decreases the ability of H-1PV to infect and kill cancer cells. This ability is rescued by the re-introduction of *LGALS1* into cancer cells. Pre-treatment with lactose, which is able to bind to galectins and modulate their cellular functions, decreased H-1PV infectivity in a dose dependent manner. In silico analysis reveals that *LGALS1* is overexpressed in various tumours including glioblastoma and pancreatic carcinoma. We show by immunohistochemistry analysis of 122 glioblastoma biopsies that galectin-1 protein levels vary between tumours, with levels in recurrent glioblastoma higher than those in primary tumours or normal tissues. We also find a direct correlation between *LGALS1* transcript levels and H-1PV oncolytic activity in 53 cancer cell lines from different tumour origins. Strikingly, the addition of purified galectin-1 sensitises poorly susceptible GBM cell lines to H-1PV killing activity by rescuing cell entry. Together, these findings demonstrate that galectin-1 is a crucial determinant of the H-1PV life cycle.

**Keywords:** oncolytic virus immunotherapy; protoparvovirus H-1PV; virus host interactions; virus cell entry; galectin-1; laminin γ1

#### **1. Introduction**

Oncolytic viruses selectively infect and destroy cancer cells while sparing normal tissues [1]. They can also stimulate strong anti-tumour immune responses and destroy tumour vasculature [2]. No fewer than 40 oncolytic viruses are currently under evaluation in clinical trials as treatments against a variety of cancers. Among them is H-1 rat protoparvovirus (H-1PV), a member of the *Parvoviridae* family in the genus *Protoparvovirus* [3,4]. This genus in addition to H-1PV includes Kilham rat virus, LuIII virus, minute virus of

**Citation:** Ferreira, T.; Kulkarni, A.; Bretscher, C.; Nazarov, P.V.; Hossain, J.A.; Ystaas, L.A.R.; Miletic, H.; Röth, R.; Niesler, B.; Marchini, A. Oncolytic H-1 Parvovirus Hijacks Galectin-1 to Enter Cancer Cells. *Viruses* **2022**, *14*, 1018. https:// doi.org/10.3390/v14051018

Academic Editor: Giorgio Gallinella

Received: 13 April 2022 Accepted: 7 May 2022 Published: 11 May 2022

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**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

mice (MVM), mouse parvovirus, tumour virus X and rat minute virus [5,6], which are also evaluated at the preclinical level as oncolytic viruses.

The H-1PV genome is a linear, single-stranded DNA molecule of around 5 kb containing the P4 and P38 promoters. The P4 promoter regulates the expression of the nonstructural (NS) gene unit encoding the NS1 and NS2 proteins. The P38 promoter controls the expression of the structural VP gene unit, which encodes the VP1 and VP2 capsid proteins and the non-structural small alternatively translated protein [4]. NS1 is the major regulator of viral DNA replication and gene transcription and is the major effector of virus oncotoxicity [7,8].

Preclinical studies in a number of cellular and animal models indicate that H-1PV can target a large variety of tumour cell lines from different tumour entities [4]. This preclinical evaluation paved the way for the clinical evaluation of H-1PV in patients with glioblastoma (GBM) [9] or pancreatic carcinoma [10]. In early-phase clinical trials, H-1PV treatment was shown to be safe and well-tolerated. Virus treatment was also associated with the first evidence of efficacy, including (i) the ability to cross the blood– brain barrier after intravenous delivery; (ii) effective distribution and expression in the tumour bed; (iii) immunoconversion of the tumour microenvironment; and (iv) improved progression-free survival and overall survival in comparison to historical controls [9]. However, treatment with H-1PV, like other oncolytic viruses, was unable to eradicate tumours in patients with the regimes used [9]. Therefore, there is an urgent need to improve the clinical outcome of H-1PV oncolytic therapy. A promising approach is the identification of host cell factors that modulate the H-1PV life cycle. This knowledge could provide hints to which drugs or treatment modalities might be combined with the virus in order to enhance its oncotoxicity. In addition, a deeper understanding of the H-1PV life cycle could help to identify biomarkers capable of predicting which patients would most likely benefit from virus treatment [11].

The first step of the virus life cycle is the virus recognition of receptor (s), co-receptor (s) or other co-factors on the cell surface modulating host cell entry. In the case of H-1PV and other protoparvoviruses, sialic acid is essential for virus–cell attachment [12–14]. Recently, we performed a druggable genome-wide siRNA library screen to identify putative modulators of H-1PV infection. The screen identified *LAMC1*, encoding the laminin γ1 chain, as a positive modulator of virus transduction. Characterisation of the interaction between H-1PV and laminin γ1 revealed that laminins, and in particular those containing laminin γ1, play a key role in mediating H-1PV attachment at the cell surface and subsequent entry into cancer cells. H-1PV binding to laminin is dependent on the sialic acid moieties in these molecules [14]. We have also shown that H-1PV cell uptake occurs through clathrin-mediated endocytosis and that the virus then passes through early to late endosomes prior to entering the nucleus. These events are dependent on dynamin activity and low endosomal pH [15].

The siRNA library screen also identified *LGALS1,* the gene encoding galectin-1 (Gal-1), as a leading activator of H-1PV infection. Interestingly, galectins are known to interact with laminins [16,17]. To date, 15 galectins have been identified in mammals [18]. They are widely expressed in various cell types and are involved in a variety of physiological functions including cell migration, mediation of cell–cell interactions, cell–matrix adhesion, transmembrane signalling, inflammation, and the immune response [19]. All galectins share a highly conserved carbohydrate-recognition domain, which binds to β-galactosides in N-linked and O-linked glycoproteins [20]. However, despite their similarities, galectins have notably different binding properties. Galectins are increasingly recognised as mediators of viral infections. However, the specific outcome of a galectin-virus interaction depends heavily on the particular galectin, the cell type, the virus, and the surrounding microenvironment. For instance, Gal-1 stabilises the binding of the human immunodeficiency virus (HIV)-1 to the host CD4 receptor on the surface of T cells by crosslinking CD4 and viral gp120 [21]. By contrast, Gal-1 inhibits Influenza A virus infection by interacting

directly with the viral envelope glycoproteins [22]. In the context of protoparvoviruses, Gal-3 promotes MVM cell uptake and infection [23,24].

In view of the potential use of H-1PV as an anti-cancer therapeutic, our goal is to characterise the early events of H-1PV infection. In this study, we demonstrate that Gal-1 plays a key role in H-1PV infection at the level of virus entry.

#### **2. Materials and Methods**

#### *2.1. Cells*

Cervical carcinoma-derived HeLa, pancreatic ductal adenocarcinoma-derived BxPC3, glioma-derived, NCH125, NCH37, U251, LN308, T98G, and A172-MG cell lines were maintained in-house [14]. The NCH125 LGALS1 KO and NCH125 CRISPR Control cell lines were established in this study (see below-*Generation of LGALS1 knockout cell line*). HeLa Control and HeLa LAMC1 KD cells were established in a previous study [14]. All cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% FBS, 100 units/mL penicillin, 100 μg/mL streptomycin and 2 mM L-glutamine (all from Gibco, Thermo Fischer Scientific, Darmstadt, Germany) in a humidified incubator at 37 ◦C and 5% CO2. All cancer cell lines were regularly tested for mycoplasma contamination using a VenorGEM OneStep Mycoplasma contamination kit (Minerva Biolabs, Berlin, Germany) and tested by a human cell authentication test (Multiplexion GmbH, Mannheim, Germany).

#### *2.2. Viruses*

Both wild-type H-1PV and recombinant H-1PV harbouring the green fluorescent protein-encoding gene (recH-1PV-EGFP) were produced, purified and titrated as previously described [25,26].

#### *2.3. siRNA-Mediated Knockdown*

Cells were seeded at a density of 4 x 10<sup>4</sup> cells/well in 24-well plates and grown in 500 μL of normal growth medium. After 24 h, cells were transfected with 10 nM siRNA using Lipofectamine RNAiMAX (Thermo Fisher Scientific, Carlsbad, CA, USA) according to the manufacturer's instructions. The following siRNAs were used for the galectins study (all purchased from Life Technologies, Paisley, UK: Silencer Select *LGALS1* siRNA (Cat. N. 4390824), Silencer *LGALS3* siRNA (Cat. N. 11332) and Silencer Select Negative Control #2 siRNA (Cat. N. 4390846). The siRNA targeting the *LAMC1* gene (Cat. N. SI00035742) and the AllStars Negative siRNA (Cat. N. SI03650318) used as a negative control were purchased from Qiagen (Hilden, Germany). After 24 h, the medium was replaced, and cells were grown for an additional 24 h to allow efficient gene silencing.

#### *2.4. Viral Transduction Assay*

Depending on the experiment, after 48 h siRNA transfection or 24 h after seeding or after pre-treatment with chemical for 30 min, cells were infected for 24 h with recH-1PV-EGFP at 0.3–0.5 TU/cell. Cells were then washed once with PBS and processed for fluorescence microscopy as described below. At least three independent experiments, each performed in duplicate, were performed for every condition.

#### *2.5. Fluorescence Microscopy*

Cells washed once with PBS were fixed with 3.7% paraformaldehyde on ice for 15 min, permeabilised with 1% Triton X-100 for 10 min and stained with 4- ,6-diamidin-2 phenylindol (DAPI). Fluorescence images of enhanced green fluorescent protein (EGFP) positive cells were acquired using a BZ-9000 fluorescence microscope (Keyence Corporation, Osaka, Japan) with a 10X objective. DAPI staining was used to visualise the cell nuclei.

#### *2.6. Lactose Pre-treatment for H-1PV Transduction Analysis*

β-lactose was purchased from Sigma-Aldrich Chemie GmbH Darmstadt, Germany (Cat. No. L-3750). Lactose stock solution was freshly prepared before treatment of the cells. HeLa cells were seeded at a density of 4 × <sup>10</sup><sup>4</sup> cells/well in 24-well plates and then pre-treated with increasing amounts (50, 100, 150, and 200 mM) of lactose for 30 min and then cells were infected with recH-1PV-EGFP for 4 h and grown for an additional 20 h. Cells were then processed as described in viral transduction assay and fluorescent microscopy sections. Numbers represent the arithmetic mean percentage of EGFP-positive cells relative to the number of EGFP-positive cells observed in untreated cells, which was arbitrarily set as 100%.

#### *2.7. Western Blotting*

Standard Western blotting was performed as described previously [15]. Immunoblotting was carried out with the following antibodies: rabbit polyclonal anti-galectin-1 (HPA000646) at 1:1000 dilution, and mouse anti-β-tubulin (T8328) (both purchased from Sigma-Aldrich, Hamburg, Germany) at 1:4000 dilution; rabbit polyclonal anti-laminin gamma 1 (PA5-36300; Thermo Fisher Scientific, Carlsbad, CA, USA) at dilution 1:1000; rabbit anti-NS1 SP8 antiserum [27] and rabbit anti-VP1/2 antiserum [28] at 1:5000 dilution. Thereafter, the membrane was incubated with horseradish peroxidase-conjugated secondary antibodies (Santa Cruz, Heidelberg, Germany) used at 1:1000 dilution.

#### *2.8. Generation of the LGALS1 Knockout Cell Line*

CRISPR/Cas9-mediated knockout of *LGALS1* in NCH125 was accomplished using galectin-1 Double Nickase Plasmid ([h]sc-400941-NIC), whereas the CRISPR/Cas9 negative control was obtained using the Control CRISPR/Cas9 Plasmid (sc-418922; both from Santa Cruz). NCH125 cells were seeded in a 6-well plate at about 70% confluency. After 24 h, 2 μg of DNA were transfected using Lipofectamine LTX (Thermo Fisher Scientific, Carlsbad, CA, USA) according to the vendor's protocol. Transfected cells were selected in normal growth medium containing 1 μg/mL puromycin (Thermo Fisher Scientific, Shanghai, China) for 72 h. Individual clones were obtained by limiting dilution. Knockout was confirmed by Western blotting.

#### *2.9. Plasmid Transfection*

To rescue *LGALS1* expression, the plasmid encoding *LGALS1* gene was used (SC118705; OriGene Technologies, Inc. Rockville, MD, USA). NCH125 Control and LGALS1 KO cells were seeded at a density of 3x105 cells/well in a 6-well plate. The next day, cells were transfected with 2.5 μg of DNA using Lipofectamine LTX or mock-transfected for 48 h.

#### *2.10. Confocal Microscopy*

Cells were seeded at a density of 3.5 × <sup>10</sup><sup>3</sup> cells/spot on spot slides and grown in 50 μL of complete cellular medium. The next day, cells were infected with wild-type H-1PV at an MOI of 500 pfu/cell in a total of 70 μL of 5% fetal bovine serum (FCS)-containing medium. At 2 h post-infection, cells were fixed with 3.7% paraformaldehyde on ice for 15 min and permeabilised with 1% Triton X-100 for 10 min. Immunostaining was carried out with the following antibodies, all used at 1:500 dilution for 1 h: mouse monoclonal anti-H-1PV capsid [29] and rabbit polyclonal anti-galectin-1 (HPA000646; Sigma-Aldrich, Darmstadt, Germany). Anti-mouse Alexa Fluor 594 IgG (A11005; Thermo Fisher Scientific, Carlsbad, CA, USA) or anti-rabbit Alexa Fluor 488 IgG (A11008; Thermo Fisher Scientific, Carlsbad, CA, USA) were used as secondary antibodies. Nuclei were stained with DAPI. Images of randomly assigned cells in the green channel (galectin-1), red channel (H-1PV), or blue channel (DAPI) were acquired with a confocal microscope (Leica TCS SP5 II, Wetzlar, Germany). Picture analysis was carried out using the LAS X Software (Leica, Wetzlar, Germany).

#### *2.11. MTT Viability Assay*

To determine cell viability after virus infection, the conversion of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) (Sigma-Aldrich Chemie GmbH, Steinheim, Germany) was measured. For this purpose, cells were seeded on a 96-well plate at a density of 2000 cells/well in 50 μL of culture medium supplemented with 10% FCS. The next day, 50 μL of serum-free medium containing wild-type H-1PV were added on top of the cells. In rescue experiments, cells were treated with H-1PV at an MOI of 5 pfu/cell, or 5 μg/mL of recombinant galectin-1 (ab50237; Abcam, Cambridge, UK), or both simultaneously. Every 24 h post-treatment, for a total of 4 time points, 10 μL of 5 mg/mL MTT were added and subsequently incubated for 2 h at 37 ◦C. Thereafter, the supernatant was aspirated, and the plates were air-dried at 37 ◦C overnight. To solubilise the formazan product, cells were then incubated with 100 μL of isopropanol for 20 min with moderate shaking and the absorbance was read with an ELISA reader at 570 nm. Viability of treated cells was expressed as a ratio of the measured absorbance (arithmetic mean of three replicates per condition) to the corresponding absorbance of untreated cells (arbitrarily defined as 100%).

#### *2.12. Binding-Only and Binding and Entry Assays*

First, the culture medium was removed and replaced with 200 μL serum-free medium containing H-1PV at an MOI of 5 pfu/cell (or H-1PV at an MOI of 5 pfu/cell and 5 μg/mL of recombinant galectin-1 simultaneously in rescue experiments). Infection was performed for 1 h at 4 ◦C to allow only cell surface virus binding or for 4 h at 37 ◦C to also allow virus cell internalisation. Thereafter, cells were extensively washed with PBS, trypsinised for 5 min, quenched with serum-containing medium and subjected to three snap freeze-thaw cycles to release cell-associated viral particles. Viral DNA was purified from cell lysates using the QiAamp MinElute Virus Spin kit (Qiagen, Hilden, Germany) according to the manufacturer's instructions. Cell-associated H-1PV genomes were quantified by following a parvovirus-specific quantitative PCR (qPCR) protocol, as previously described [12]. A minimum of three independent experiments were performed in triplicate for every condition tested.

#### *2.13. Flow Cytometry*

Cells were seeded at a density of 5 × <sup>10</sup><sup>5</sup> cells/well in a 6-well plate. The next day, cells were infected with H-1PV at an MOI of 25 pfu/cell for 1 h at 4 ◦C. Cells were washed with ice-cold PBS and then gently scraped off with a cell lifter on ice. Cells were then fixed with 2% formaldehyde for 15 min at 4 ◦C and blocked with 2.5% albumin bovine fraction V (BSA; SERVA Electrophoresis, Heidelberg, Germany)/PBS for 20 min at 4 ◦C. Cells were then incubated with H-1PV anti-capsid antibody (dilution 1:500) for 30 min at 4 ◦C, and subsequently with Alexa Fluor 488 goat α-mouse (1:500) for 30 min at 4 ◦C. Three washes with 2.5% BSA/PBS were performed between each staining step. Analysis was carried out using a FACS Calibur (BD Biosciences, San Jose, CA, USA).

#### *2.14. Tissue Microarray*

Patient material for the tissue microarray was derived from paraffin embedded GBM biopsies obtained from the Department of Pathology, Haukeland University Hospital, Bergen, Norway. The project (number 2017/2505) was approved by the Regional Ethics Committee (Bergen, Norway). Control tissues (brain, liver and tonsil) were derived from autopsy material. The microarray included 61 primary GBM and 49 recurrent GBM biopsies, plus 12 biopsies from normal tissues (four each from brain, tonsil and liver). Immunohistochemical staining was carried out as described previously [30] using galectin-1 antibody (sc-166618; Santa Cruz, Santa Cruz, CA, USA) at a dilution of 1:200 followed by a biotinylated anti-mouse antibody (Vector Laboratories, Burlingame, CA, USA) at a dilution of 1:1100. Galectin-1-positive cells were counted via automated counting, as described previously [31].

#### *2.15. Correlation Analysis between Gene Expression of Cancer Cell Lines and H-1PV-Induced Oncolysis*

Gene expression data were taken from two databases: 53 cancer cell lines of NCI-60 dataset (https://discover.nci.nih.gov/cellminer, accessed on 23 January 2021, RNAseq data) and the Cancer Cell Line Encyclopedia (CCLE, 52 cancer cell lines) [32]. Simple linear regression models were built for 2 considered genes of interest (*LGALS1* and *LAMC1*) and 2 control genes (*LGALS3* and *GAPDH*), predicting experimentally observed EC50 values. Additionally, a two-variable regression was built predicting EC50 with both genes *LGALS3* and *GAPDH*. Significance of the models was characterised by the *p*-values (*p*), coefficients of determination (*R*2) and Pearson correlations (*R*).

#### *2.16. Measurement of Transcript Levels*

The mRNA expression levels of the target of interest *LGALS1*, as well as of the reference genes *ACTB*, *GAPDH* and *PGK1*, were quantified at the nCounter Core Facility on a SPRINT Profiler system by nCounter technology (NanoString Technologies (Seattle, WA, USA), as described previously [14]. Accession numbers and target sequences of analysed genes are:

*LGALS1* gene (accession number NM\_002305.4): GGTGCGCCTGCCCGGGAACATC-CTCCTGGACTCAATCATGGCTTGTGGTCTGGTCGCCAGCAACCTGAATCTCAAACCT GGAGAGTGCCTTCGAGTGCGA

*ACTB* gene (accession number NM\_001101.2): TGCAGAAGGAGATCACTGCCCTG-GCACCCAGCACAATGAAGATCAAGATCATTGCTCCTCCTGAGCGCAAGTACTCCGT GTGGATCGGCGGCTCCATCCT

*GAPDH* gene (NM\_001256799.1): GAACGGGAAGCTTGTCATCAATGGAAATCC-CATCACCATCTTCCAGGAGCGAGATCCCTCCAAAATCAAGTGGGGCGATGCTGGCG CTGAGTACGTCGTG

*PGK1* gene (NM\_000291.2): GCAAGAAGTATGCTGAGGCTGTCACTCGGGCTAAGC AGATTGTGTGGAATGGTCCTGTGGGGGTATTTGAATGGGAAGCTTTTGCCCGGGGAA CCAAAGC

#### *2.17. xCELLigence*

Cell proliferation was monitored in real time through the xCelligence system (ACEA Biosciences Inc. San Diego, CA, USA) according to the manufacturer's instructions. Briefly, <sup>8</sup> × 104 cells per well were seeded in a 96-well E-plate (Roche, Mannheim, Germany) in a total volume of 100 μL of complete DMEM medium. Cells were treated during the cellular growth phase with H-1PV at an MOI of 5 pfu/cell or 5 μg/mL of recombinant galectin-1, or both simultaneously. Cell proliferation was monitored every 30 min in real time. Data are expressed as "Cell index" (*n* = 3) calculated by the RTCA software 1.2.1 (Agilent) as a measure of cell adhesion and, therefore, cell viability.

#### *2.18. Statistical Analysis*

Results are shown as the arithmetic mean of biological replicates ± standard deviation (SD) from a representative experiment. Statistical significance was determined by a paired two-tailed Student's *t*-test, unless stated otherwise, using Microsoft Excel 365 and/or GraphPad Prism version 8. Only values below 0.05 were considered significant: *p* ≤ 0.05 (\*), *p* ≤ 0.01 (\*\*) and *p* ≤ 0.001 (\*\*\*).

#### **3. Results**

#### *3.1. Knockdown of LGALS1, but Not LGALS3, Hampers H-1PV Infection*

To identify cellular modulators of the H-1PV life cycle, we previously carried out a high-throughput siRNA library screening in cervical carcinoma-derived HeLa cells using a siRNA library targeting the human druggable genome (6961 genes, each targeted by a pool of four siRNAs/gene) [14]. This led to the identification of laminins, in particular those containing the laminin γ1 chain, as factors used by H-1PV to attach at the cell surface and to enter cancer cells [14]. In the same screening, *LGALS1*—the gene encoding Gal-1emerged as another top activator of H-1PV transduction, as its silencing decreased H-1PV transduction by approximately 70% [14]. These findings, along with the fact that galectins interact with laminins [17], prompted us to hypothesise that Gal-1 is involved in H-1PV infection at the level of cell entry. This hypothesis was also supported by the discovery that MVM (a closely related protoparvovirus) requires Gal-3 to efficiently infect mouse cells [23,24].

We confirmed the results of the siRNA library screening by performing a knockdown of *LGALS1* using an independent siRNA. Given the role of Gal-3 in host cell entry by MVM, we also investigated the effect of siRNA-mediated silencing of *LGALS3*. A recombinant H-1PV expressing the *EGFP* reporter gene (recH-1PV-EGFP) was used for these experiments [25]. This non-replicative parvovirus shares the same capsid of the wild type but harbours the *EGFP* gene under the control of the natural P38 late promoter, whose activity is regulated by the NS1 viral protein. Therefore, the EGFP signal directly correlates with the ability of the virus to reach the nucleus and initiate its own gene transcription.

Cervical cancer-derived HeLa, glioma-derived NCH125 and pancreatic carcinomaderived BxPC3 cell lines were transfected with siRNAs targeting *LGALS1* or *LGALS3*, or a scrambled siRNA. After 48 h, cells were infected with recH-1PV-EGFP and grown for a further 24 h. Efficient gene silencing was achieved for both genes in cells transfected with their respective siRNAs (Figure S1). However, only the siRNA targeting *LGALS1* significantly decreased H-1PV transduction (by more than 55%). As opposed to the role of Gal-3 in MVM infection, silencing of *LGALS3* did not significantly alter H-1PV transduction when compared with scramble controls (Figure 1). These results confirm the results of our siRNA library screening, which indicated that Gal-1, but not Gal-3, plays a key role in H-1PV infection in the cell lines tested.

#### *3.2. Pre-Treatment with Lactose Inhibits H-1PV Infection*

Gal-1 is a member of the galectins which belongs to a sub-family of lectins, defined by their highly conserved carbohydrate recognition domain (CRD) with the ability to bind to a number of beta-galactosides. Among the beta-galactosides, lactose is known to modulate the function of Gal-1 by directly binding to the CRD [33]. Hence, we hypothesised that treatment with lactose may interfere with H-1PV infectivity by competing with the virus for the interaction with Gal-1. To test this hypothesis, HeLa cells were pre-treated with increasing concentrations of soluble β-lactose before being infected with recH-1PV-EGFP. Pre-treatment with lactose decreased H-1PV transduction in a dose dependent manner (Figure 2). These results provide further evidence that H-1PV interaction with Gal-1 plays a crucial role in modulating H-1PV infectivity.

**Figure 1.** H-1PV transduction is reduced in *LGALS1*, but not *LGALS3*, knockdown cell lines. HeLa, NCH125 and BxPC3 cells were transfected with siRNAs targeting *LGALS1* or *LGALS3* or with a scrambled siRNA. At 48 h post-transfection, cells were infected with recH-1PV-EGFP for 4 h and grown for an additional 20 h. Cells were then processed as described in the Materials and Methods.

Numbers represent the arithmetic mean percentage of EGFP-positive cells relative to the number of EGFP-positive cells observed in cells transfected with control siRNA, which was arbitrarily set as 100%. The independent experiment shown was repeated thrice each with three biologically independent samples (*ns–not significant*; \*\*\* *p* ≤ 0.001, calculated by using a one-way ANOVA).

**Figure 2.** Pre-treatment with lactose decreases H-1PV infection in a dose-dependent manner. HeLa cells were pre-treated with the indicated concentrations of β-lactose for 30 min and then infected with recH-1PV-EGFP for 4 h. At 20 h after infection, cells were then harvested and processed as described in the Materials and Methods section. Numbers represent the arithmetic mean percentage of EGFP-positive cells relative to the number of EGFP-positive cells observed in untreated cells, which was arbitrarily set as 100%. The independent experiment shown was repeated thrice each with three biologically independent samples (*ns–not significant; \*\* p* > 0.05; \*\*\* *p* ≤ 0.001, calculated by using a one-way ANOVA).

#### *3.3. Galectin-1 Knockout Impairs H-1PV Infection in NCH125 Cells*

To further investigate the biological role of Gal-1 in H-1PV infection, we took advantage of the CRISPR-Cas9 technology and established the NCH125 *LGALS1* knockout cell line (LGALS1 KO). We also established the NCH125 control cell line (Control) using a non-targeting guide RNA control sequence. Given that H-1PV requires S-phase factors expressed in proliferating cells for a productive infection [8], we evaluated the proliferation of LGALS1 KO versus Control cells. Both cell lines proliferated at a similar rate, as shown by real-time monitoring of cell growth and viability via xCELLigence (Figure S2).

We infected LGALS1 KO and Control cell lines with wild-type H-1PV and analysed the number of internalised virus particles by immunofluorescence using a specific anti-capsid antibody [29]. Two hours post-infection, fluorescence was significantly lower in LGALS1 KO cells than in Control cells (Figure 3A). Then, we evaluated the levels of viral proteins 48 h post-infection with the wild-type H-1PV. In agreement with previous results, Western blotting analysis revealed that NS1 and VP1 protein levels were lower in LGALS1 KO cells than in Control cells (Figure 3B). Finally, we analysed H-1PV transduction efficiency by infecting both cell lines with recH-1PV-EGFP. Consistent with reduced H-1PV entry, a significant decrease in transduction activity was found in LGALS1 KO cells (47%) in comparison with Control cells (Figure 3C). Strikingly, transient transfection of LGALS1 KO cells with a plasmid encoding *LGALS1* 48 h prior to infection, by re-establishing Gal-1 protein levels to values similar to those observed in Control cells (Figure 3C: Western blotting analysis), rescued the reduction in H-1PV transduction in these cells (Figure 3C).

**Figure 3.** H-1PV infectivity is reduced in NCH125 LGALS1 KO cells. (**A**) H-1PV entry decreases in NCH125 LGALS1 KO cells. Control and LGALS1 KO cells were infected with H-1PV at an MOI of 50 pfu/cell for 2 h at 37 ◦C and prepared for confocal microscopy analysis. Gal-1 (green) and H-1PV capsid (red) were detected using specific antibodies, while DAPI (blue) was used to stain nuclei. As expected, Gal-1 was readily detected in Control cells, while it fell below detection limits in LGALS1 KO cells. The lower panel shows representative examples of H-1PV-infected cells. Quantification of the H-1PV fluorescence signal is shown on the right. This was retrieved from two independent experiments in which the fluorescence intensity was quantified in 25 randomly identified cells using ImageJ. Box plot depicts the median with a centre line, and the Tukey–Whiskers plots indicate variability outside the upper and lower quartiles (*n* = 25, \*\*\* *p* ≤ 0.001). (**B**) Control and LGALS1 KO cells were infected with H-1PV at an MOI of 2 pfu/cell for 48 h, and NS1 and VP1 protein levels were assessed by Western blotting. Beta-tubulin was used as a loading control. (**C**) H-1PV transduction is decreased in LGALS1 KO cells and re-established by transfecting the cells with a plasmid carrying *LGALS1*. LGALS1 KO cells were transfected with a plasmid encoding *LGALS1*, treated only with lipofectamine LTX (mock transfection) or left untreated. In addition, 48 h post-transfection, cells were infected with recH-1PV-EGFP for 24 h. Control cells were also included, and the level of virus transduction was set arbitrarily at 100%. The independent experiment shown was repeated twice each with four biologically independent samples (*ns*: *p* > 0.05; \*\*\* *p* ≤ 0.001, calculating using a one-way ANOVA). On the right side, Western blotting analysis shows the levels of Gal-1 at the time of infection. Beta-tubulin was used as a loading control.

#### *3.4. Galectin-1 Knockout Decreases H-1PV Oncolytic Activity in NCH125 Cells*

As *LGALS1* knockdown/out decreased the overall amount of internalised H-1PV, we assessed whether this would result in reduced oncolytic activity. For this purpose, we assessed the susceptibility of LGALS1 KO and Control cell lines to H-1PV infection in a time course experiment in which viability of infected cells was assessed every 24 h for a total of 96 h. Whereas the viability of Control cells decreased progressively over time, the viability of LGALS1 KO cells, which were less sensitive to H-1PV infection, remained high throughout the experiment (above 75%; Figure 4A). Remarkably, the susceptibility of LGALS1 KO cells to H-1PV oncotoxicity was re-established by infecting the cells together with human recombinant Gal-1. Indeed, the viability of LGALS1 KO cells infected with H-1PV dropped from 74% to 18% in the presence of Gal-1; a control experiment showed that the protein itself was not toxic to the cells at the concentrations used (Figure 4B). Together, these results highlight the critical role of Gal-1 in H-1PV infection in NCH125 cells.

#### *3.5. Galectin-1 Plays a Role in H-1PV Virus Entry Rather Than Cell Surface Attachment*

The results shown above indicate that Gal-1 is involved in the early steps of H-1PV infection. However, whether Gal-1 is required for H-1PV attachment at the cell surface, internalisation, or both events, remains to be determined. To elucidate the role of Gal-1 in H-1PV infection, we first performed virus binding and entry assays. LGALS1 KO and Control cell lines were infected with wild-type H-1PV at 37 ◦C for different times (0.5, 1, 2 and 4 h) and then cell-associated viral DNA was quantified by PCR. In agreement with previous results, we observed less cell-associated H-1PV DNA in LGALS1 KO cells than in Control cells (Figure 5A). The addition of recombinant Gal-1 increased the number of cell-associated H-1PV genomes in LGALS1 KO cells to values that were similar to those found in H-1PV-infected Control cells (Figure 5B). These results confirm the role of Gal-1 in H-1PV binding and/or entry.

**Figure 4.** H-1PV has reduced oncolytic activity in NCH125 LGALS1 KO cells, which is rescued by supplementing with recombinant Gal-1 protein; (**A**) H-1PV oncolytic activity is reduced in NCH125 LGALS1 KO cells. Control and LGASL1 KO cells were infected with H-1PV at an MOI of 1 pfu/cell. Cell viability was assessed by MTT every 24 h for a total of 96 h. The curve plot depicts the mean ± standard deviation for each time point expressed as a percentage of cell viability compared to corresponding uninfected cells. The independent experiment shown was repeated thrice each with four biologically independent samples (\*\*\* *p* ≤ 0.001). (**B**) Purified Gal-1 rescues H-1PV oncolytic activity in NCH125 LGALS1 KO cells. Control and LGASL1 KO cells were infected (or not) with H-1PV at an MOI of 1 pfu/cell, in the presence or absence of 5 μg/mL of human recombinant Gal-1. Cell viability was assessed at 72 h post-infection by MTT. Columns depict the percentage (mean value) of cell viability compared to uninfected cells ± standard deviation bars. The independent experiment shown was repeated twice each with four biologically independent samples (*ns: p* > 0.05; \*\*\* *p* ≤ 0.001, calculated using a one-way ANOVA).

Concerning the possible involvement of Gal-1 in H-1PV cell surface attachment, LGALS1 KO and Control cell lines were inoculated with H-1PV at 4 ◦C for 1 h. Under these conditions, only virus attachment at the cell surface virus occurs, while cell entry is prevented [14]. After removing unbound H-1PV particles, those that remained attached to the cell surface were stained with an anti-capsid antibody and Flow cytometry analysis was performed. No significant differences in the fluorescence signal (H-1PV binding) were observed between the LGALS1 KO and Control cells (Figure 5C). The same findings were obtained by quantitative PCR analysis (Figure 5D). These results speak for an involvement of Gal-1 in H-1PV entry rather than in H-1PV binding to the cell surface.

**Figure 5.** H-1PV cell entry, but not cell attachment, is reduced in NCH125 LGALS1 KO cells. (**A**) H-1PV binding and entry assay assessed by qPCR. NCH125 Control and LGALS1 KO cells were infected with H-1PV at an MOI of 5 pfu/cell for 0.5, 1, 1.5, 2 and 4 h at 37 ◦C. Cells were then extensively washed and harvested, and encapsidated viral DNA was extracted and subjected to qPCR. Columns in the graph show the number of copies of the cell-associated H-1PV genome with relative standard deviations (\*\*\* *p* ≤ 0.001). The independent experiment shown was repeated thrice; *n* = 3 biologically independent samples. (**B**) Binding and entry is rescued by the addition of purified recombinant Gal-1. At the time of H-1PV infection (at an MOI of 5 pfu/cell), Gal-1 was added (or not) to the culture medium. Infection was carried out for 4 h. Numbers indicate the percentage of cell-associated genomes relative to NCH125 Control cells infected with H-1PV arbitrarily set as 100% The independent experiment shown was repeated twice each with four biologically independent samples (*ns: p >* 0.05; \*\*\* *p* ≤ 0.001, calculated using a one-way ANOVA). (**C**) H-1PV cell surface binding assessed by Flow cytometry. A representative flow cytometry histogram with overlay of Control (black) and LGALS1 KO cells (blue) shows no difference in H-1PV-associated cells. Cells were

either mock- or H-1PV-infected (at an MOI of 25 pfu/cell) for 1 h at 4 ◦C. Cells were not permeabilised for the Flow cytometry analysis, and cell surface-bound H-1PV particles were detected with a specific anti-capsid antibody. The independent experiment shown was repeated twice each with two biologically independent samples. (**D**) H-1PV binding only, assessed by qPCR. Control and LGALS1 KO cells were infected with H-1PV (at an MOI of 5 pfu/cell) for 1 h at 4 ◦C. Cells were then washed and harvested, and extracted encapsidated viral DNA was quantified by qPCR. The independent experiment shown was repeated thrice each with three biologically independent samples.

#### *3.6. Gal-1 Cooperates with Laminins in Mediating H-1PV Infection*

Given that laminins containing the γ1 chain play determinant roles in H-1PV attachment at the cell surface as well as in H-1PV cell entry, we asked whether siRNAmediated silencing of *LAMC1* in LGALS1 KO cells would further decrease H-1PV infection (Figure S3). In agreement with the results shown above, reduced H-1PV cell uptake was observed in LGALS1 KO cells. Furthermore, given the important role of laminins in H-1PV cell attachment and entry, knockdown of *LAMC1* gene expression also strongly reduced cell-associated H-1PV genomes in NCH125 cells. The reduction observed upon *LAMC1* knockdown was not significantly different between LGALS1 KO cells and Control cells, indicating that, in the absence of Gal-1, the depletion of *LAMC1* does not further reduce H-1PV cell uptake (Figure 6A). To confirm these results, we repeated the double knock-down experiment in HeLa cells (Figure S3). To this end, we silenced *LGALS1* gene expression in the previously established HeLa LAMC1 KD cell line (in which the *LAMC1* gene was knocked down via CRISPR-Cas9) [14]. In agreement with published results, infection of HeLa LAMC1 KD cells presented a 40% reduction in cell-associated H-1PV genomes in comparison with Control cells. The silencing of *LGALS1* also decreased H-1PV cell uptake in HeLa cells, yet no significant difference was observed between HeLa control and HeLa LAMC1 KD cell lines (Figure 6B). The fact that removal/reduction of both laminin γ1 and Gal-1 does not synergistically inhibit infection suggests that the two factors may act on the same H-1PV entry pathway. At the same time, the finding that, under these conditions, a fraction of viral particles is still able to penetrate cells supports the idea that H-1PV may also use alternative pathways and exploit other cellular factors to infect cells.

#### *3.7. Gal-1 Is a Marker of Bad Prognosis in Various Tumour Types including GBM*

Growing evidence indicates that overexpression of *LGALS1* is associated with metastasis formation, tumour recurrence and poor tumour prognosis [34]. Our analysis of brain tumour expression datasets using the GlioVis web application (http://gliovis.bioinfo.cnio.es/, accessed on 23 January 2021) revealed that *LGALS1* overexpression is associated with worse overall survival for brain tumours. Focusing particularly on GBM, we observed that these tissues have significantly higher expression of *LGALS1* in comparison to those from healthy individuals (Figure S4A) and *LGALS1* levels increase with the severity of the malignancy from WHO grade II to IV (Figure S4B). High *LGALS1* expression is associated with poor prognosis in glioma (Figure S5).

Next, we investigated whether Gal-1 protein levels varied between normal tissues, primary and recurrent GBM tissues. To this end, we used an in-house protein tissue microarray including a cohort of 110 GBM patient biopsies (61 primary and 49 recurrent GBM) and 12 biopsies from normal tissues (four each from brain, liver, and tonsil) and performed immunohistochemistry using an anti-galectin-1 antibody. Levels of Gal-1 were higher overall in GBM biopsies than in normal tissues. Among the GBM biopsies, we found a diversified Gal-1 expression profile with recurrent GBM tissues expressing significantly higher levels of Gal-1 than primary GBM tissues (45% of recurrent GBM tissues expressed medium or high levels of Gal-1, compared with 20% of primary GBM tissues; Figure 7).

**Figure 6.** Effect on H-1PV binding and entry upon depletion of both *LAMC1* and *LGALS1* in NCH125 and HeLa cells. (**A**) NCH125 Control and LGALS1 KO cells were transfected with a siRNA targeting *LAMC1* or a negative control siRNA. At 48 h post-transfection, cells were infected with H-1PV at an MOI of 5 pfu/cell for 4 h at 37 ◦C. Cells were then extensively washed and harvested, and encapsidated viral DNA was extracted and subjected to qPCR. Columns in the graph show the number of copies of the cell-associated H-1PV genome with relative standard deviations (*ns*: *p* > 0.05; \*\*\* *p* ≤ 0.001, calculated using a one-way ANOVA). The independent experiment shown was repeated thrice each with threebiologically independent samples. (**B**) HeLa Control and HeLa LAMC1 KD cells were transfected with a siRNA targeting *LGALS1* or a negative control siRNA. After 48 h siRNA transfection, cells were infected with H-1PV at an MOI of 5 pfu/cell for 4 h at 37 ◦C. Cells were then processed as per **A**. The experiment was performed with four biologically independent samples (\*\* *p* ≤ 0.01, calculated using a one-way ANOVA).

#### *3.8. LGALS1 Expression Profile of NCI-60 Cells Positively Correlates with H-1PV Oncotoxicity*

Given the important role of Gal-1 in H-1PV infection, we looked for a putative correlation between *LGALS1* expression levels and H-1PV oncotoxicity. To this end, we screened 53 cancer cell lines from the NCI-60 panel for their susceptibility to H-1PV [14]. The gene expression profiles of these cell lines are fully characterised and publicly available [35]. We assessed virus-mediated oncotoxicity by monitoring cell viability in real time using xCELLigence [14]. We calculated the viral MOI responsible for killing 50% of the cell population at 72 h post-infection (EC50). Using gene expression data from NCI-60 and the Cancer Cell Line Encyclopedia (CCLE), we found that *LGALS1* mRNA expression levels anti-correlated with EC50 values, suggesting that cells expressing higher levels of *LGALS1* may be more susceptible to virus killing activity (Figure 8).

**Figure 7.** Differential expression of Gal-1 in normal tissues and in primary and recurrent GBM biopsies. (**A**) overview of the tissue microarray. This study included biopsies from normal tissue (*n* = 12), primary GBM biopsies (*n* = 61) and recurrent GBM biopsies (*n* = 49). Biopsies were categorised based on Gal-1 expression after immunostaining with anti-galectin-1 antibody: low (<10% positive cells), medium (10–30% positive cells) or high expression (>30% positive cells). The number of biopsies in each category are indicated under each representative image. The staining was performed twice in each normal sample and thrice on tumour tissues. Quantification of Gal-1-positive cells (%) was performed as described in the Materials & Methods section using in-house software; (**B**) comparative analysis of Gal-1 expression between healthy tissues and GBM biopsies (primary and recurrent); (**C**) comparative analysis of Gal-1 expression between primary and recurrent GBM biopsies. The arithmetic mean of Gal-1-positive cells is indicated with a horizontal line and by the number above (\*\* *p*≤ 0.01; \*\*\* *p* ≤ 0.001).

**Figure 8.** Correlation between *LGALS1* gene expression of cancer cell lines and their susceptibility to H-1PV-induced oncolysis. (**A**) *LGALS1* gene expression was retrieved from the National Cancer Institute (NCI)-60 database. Fifty-three cancer cell lines from the NCI-60 panel were tested for their susceptibility to H-1PV infection by xCELLigence. H-1PV EC50 values were calculated as the viral MOI that kills 50% of the cell population at 72 h post-infection (72hpi), measured by xCELLigence (see also [14]). Six cancer cell lines (MCF7, COLO 205, HCC-2998, HCT-15, LOX IMVI, OVCAR-3 (indicated by arrows) were found to be resistant to cell lysis even at the maximum tested concentrations of H-1PV (MOI 50 pfu/cell). Therefore, as EC50 values could not be calculated for those cell lines, their values were arbitrarily fixed as 100; (**B**) *LGALS1* expression versus H-1PV EC50. Each blue dot corresponds to a cell line and the grey line corresponds to a linear regression. (**C**) *LGALS1* levels are moderately anti-correlated with H-1PV EC50. *LGALS1* gene expression measurements were retrieved from the NCI-60 (53 cell lines) and Cancer Cell Line Encyclopedia (CCLE) (52 cell lines). Bar plot depicts the correlation between the gene expression from each dataset and the EC50 values (Pearson's correlation). Significant anti-correlation was observed for both NCI-60 and CCLE datasets with *R* = –0.510, C.I. (–0.685, –0.277), *p* < 0.001 and –0.422, C.I. (–0.641, –0.139), *p* < 0.01 respectively (in both cases, the null hypothesis: *R* = 0).

A similar anti-correlation was previously shown for *LAMC1* [14]. Therefore, we asked whether we could better predict the susceptibility of a certain cancer cell to H-1PV infectivity and cell killing by analysing the expression levels of the two genes together. As negative controls, *LGALS3* (selected for showing no apparent role in H-1PV infection) and *GAPDH* were chosen. Using both NCI-60 and the CCLE databases, we found that a slight increase in predictability of EC50 may be achieved by combining *LAMC1* and *LGALS1* in a single linear regression model. For NCI-60, *R*<sup>2</sup> of the combined model was 0.313 (*<sup>p</sup>* = 8.3 × <sup>10</sup><sup>−</sup>5), while simple models gave *<sup>R</sup>*<sup>2</sup> of 0.188 (*LAMC1*, *<sup>p</sup>* = 1.2 × <sup>10</sup><sup>−</sup>3, *<sup>R</sup>* <sup>=</sup> −0.433) and 0.260 (*LGALS1*, *<sup>p</sup>* = 9.8 × <sup>10</sup>−5, *<sup>R</sup>* = –0.510). Control genes show no significant linear relation (*LGALS3*: *R*<sup>2</sup> = 0.015, *p* = 0.38, *GAPDH*: *R*<sup>2</sup> = 0.018, *p* = 0.34) (Figure S6). Similar results were obtained using the CCLE dataset (Figure S7). These results are in line with the important role that laminin γ1 and Gal-1 play in H-1PV cell surface recognition and entry.

#### *3.9. LGALS1 Expression Positively Correlates with H-1PV Oncolysis in Glioma Cell Lines*

Glioma cancer cell lines are generally susceptible to H-1PV oncolysis [36]. However, not all cancer cell lines respond similarly to H-1PV oncolysis: they range from highly to lowly permissive, or are even resistant. We recently described four glioma cell lines that are semi-permissive to H-1PV infection, namely U251, LN308, T98G and A172-MG, which all express low levels of *LAMC1* mRNA [14]. However, it is possible that other cell components may account for the poor susceptibility of these cell lines to virus infection. Using the NanoString technology, we found that *LGALS1* mRNA levels were lower in the four aforementioned cell lines, as well as in the control normal human astrocytes, than in two H-1PV-sensitive glioma cell lines (NCH125 and NCH37) (Figure 9A). Monitoring of cell viability in real time by xCELLigence confirmed our previous results showing that NCH125 and NCH37 cell lines are efficiently killed by H-1PV at an MOI of 5 (pfu/cell). By contrast, U251, LN308, T98G, A172-MG cell lines as well as control normal human astrocytes were not. Remarkably, susceptibility of the four semi-permissive glioma cell lines to H-1PV oncotoxicity was substantially enhanced by the addition of human recombinant Gal-1 (Figure 9B). Consistent with our previous results, the addition of exogenous Gal-1 promoted H-1PV entry in the four glioma cell lines leading to an increase in the number of cell-associated viral genomes by 1.5- to 2.8-fold (Figure 9D), while not interfering with viral binding to the cell surface (Figure 9C). Together, these results confirm that Gal-1 plays a critical role in H-1PV infection at the level of virus entry and provide evidence that Gal-1 levels can determine the outcome of H-1PV infection. These results further support the importance of Gal-1 to H-1PV oncolytic activity and pave the way for its use in predicting the success of H-1PV infection.

**Figure 9.** Galectin-1 levels in glioma cell lines determine the success of H-1PV infection. (**A**) Total mRNA was isolated from glioma cell lines susceptible (NCH125; NCH37) or semi-permissive (U251; LN308; T98G; A172-MG) to H-1PV infection, and *LGALS1* mRNA transcripts were measured using nCounter analysis. Bar graph depicts the *LGALS1* transcript counts; numbers on the top of the columns indicate gene expression fold changes between susceptible and semi-permissive cancer cell lines. The independent experiment is shown; *n* = 1 (NCH125, NCH37, U251 and A172-MG); *n* = 2 (LN308 and T98G); *n* = 3 (human astrocytes) biologically independent samples. (**B**) NCH125 and NCH37 cell lines were either infected with H-1PV at an MOI of 5 pfu/cell (green) or left untreated (red). Semi-permissive cell lines were infected with H-1PV at an MOI of 5 pfu/cell (green), incubated with 5 μg/mL of human recombinant Gal-1 (pink), H-1PV and Gal-1 simultaneously (blue), or left untreated (red). Cell viability was assessed by xCELLigence every 30 min in real time. The curve shows the "Cell index" mean of three biologically independent samples (*n* = 3) at any given time, which is proportional to the viability of the cell population. Black arrows indicate the time of treatment. (**C**) H-1PV binding at the cell surface is not affected by Gal-1 addition. U251, LN308, T98G

and A172-MG cells were incubated with H-1PV alone at an MOI of 5 pfu/cell or with H-1PV and Gal-1. Incubations were carried out for 1 h at 4 ◦C (binding only). Cells were then washed and harvested, and encapsidated viral DNA was extracted and subsequently quantified by qPCR. Columns in the graph show the fold change of number of copies of cell-associated H-1PV genome relative to the virus-infected cells arbitrarily set as 1, with respective standard deviations. The independent experiment shown was performed with four biologically independent samples (*ns*: *p* > 0.05; \*\*\* *p* ≤ 0.001, calculated using a one-way ANOVA). (**D**) H-1PV entry is rescued upon Gal-1 addition. U251, LN308, T98G and A172-MG cells were incubated with H-1PV alone at an MOI of 5 pfu/cell or with H-1PV and Gal-1. Incubations were carried out either for 4 h at 37 ◦C (binding and entry). Cells were processed and results analysed as described in C.

#### **4. Discussion**

Following more than five decades of preclinical research, H-1PV monotherapy has been evaluated in patients with recurrent GBM and pancreatic carcinoma in early-phase clinical trials. H-1PV treatment was demonstrated to be safe, well tolerated and associated with surrogate evidence of anticancer efficacy, including immunoconversion of the tumour microenvironment and improved progression-free survival and overall survival in comparison with historical controls [9]. These promising results have motivated research aiming at further improving H-1PV efficacy [11,37].

The virus life cycle is a multistep process that is heavily dependent on the presence and abundance of viral (co-)receptors, processing enzymes and proteins required for a productive infection. The levels and activities of these components may vary in different cancer cells, determining their susceptibility to a particular virus. We anticipate that a better understanding of the H-1PV life cycle may provide the cues to further develop H-1PV-based therapies. For instance, this knowledge may help to identify new drugs that enhance H-1PV replication in cancer cells and/or oncolytic activity by modulating H-1PV-related cellular pathways. Furthermore, the cellular factors involved in the H-1PV life cycle may also serve as biomarkers to predict whether a certain tumour is susceptible or resistant to H-1PV infection.

Recently, we found that H-1PV enters cancer cells via clathrin-mediated endocytosis, a process that involves dynamin and requires a low pH in the endocytic compartments [15]. We also reported that laminins, in particular those containing the laminin γ1 chain, act as attachment factors at the cell surface for a successful H-1PV infection [14]. In particular, we found that sialic acid moieties in the laminins provide a docking place for the virus to anchor to at the cell surface [14]. Laminin γ1 was originally identified as a modulator of the H-1PV life cycle in a siRNA screening using a druggable-genome library performed in HeLa cells. The same screening revealed Gal-1 as another top activator of H-1PV infection. Indeed, silencing of *LGALS1* impaired H-1PV virus transduction by approximately 70% in HeLa cells. These results prompted us to explore whether Gal-1 is involved in the early steps of H-1PV infection.

In the present study, we show that Gal-1 plays a central role in H-1PV infection at the level of H-1PV cell entry but not cell attachment, indicating a role that is distinct from that of laminins in the virus cell cycle. By contrast, knockdown of *LGALS3* did not impair H-1PV infection (Figure 1), further supporting the specificity of the interaction between H-1PV and Gal-1.

Until the present study, Gal-3 was the only galectin that had been implicated in a protoparvovirus infection. Indeed, knockdown of *LGALS3* rendered LA9 mouse fibroblasts less susceptible to MVM infection. This phenotype was not due to reduced binding to the cell surface; instead, Gal-3 was responsible for promoting efficient virus uptake [24]. Our results indicate that Gal-1 is essential for a productive H-1PV infection at the level of cell entry, with no evidence of its requirement in viral binding to the plasma membrane in NCH125 cell lines. Therefore, these findings suggest that the mechanisms through which Gal-1 mediates H-1PV entry are similar to those of Gal-3 in MVM infection [23]. The fact that our results do not support an involvement of Gal-3 or clathrin-independent endocytosis in the H-1PV entry process [15] suggests that H-1PV and MVM engage different galectins for their entry processes which may contribute to their different tropism. At this point, we cannot also exclude that, in addition to Gal-1, other members of the galectin family may participate in the H-1PV entry process.

Based on our results, we envision a model in which H-1PV interacts with different classes of molecules, rather than with a single cell surface receptor, in order to enter cancer cells. Laminins containing γ1 chains would accumulate virus in the vicinity of the cell surface via sialic acid, while Gal-1 would promote the efficient internalisation of virus particles into a clathrin-coated pit. After engagement of these factors, H-1PV would penetrate the cells preferentially via clathrin-mediated endocytosis [15].

One possibility arises that H-1PV hijacks extracellular Gal-1 to enter the cells together with the protein. Our results support this idea by showing that the addition of exogenous purified Gal-1 boosts H-1PV infection at the level of virus entry, thereby sensitising semipermissive cancer cells to H-1PV-mediated oncolysis. On the other hand, we cannot exclude that Gal-1 may play additional roles at post-entry levels, including viral trafficking and DNA uncoating.

Galectins are known to be synthesised in the cytoplasm and accumulate there until they are secreted via a poorly characterised pathway [38]. The exact mechanism through which galectin(s) translocate across the cell membrane remains also poorly understood [39]. However, previous research has shown that inhibition of the lipid raft-dependent pathway does not impede Gal-1 internalisation; instead, a total block of Gal-1 internalisation was observed only when both clathrin-mediated endocytosis and lipid rafts were disrupted, demonstrating that Gal-1 enters cells through various mechanisms, including clathrinmediated endocytosis [40,41]. Therefore, it may be possible that H-1PV uses Gal-1 to enter cancer cells through clathrin-mediated endocytosis.

Alternatively, Gal-1 could mediate the binding of H-1PV to other cellular factors involved in its entry, e.g. a transmembrane receptor or a co-receptor. A number of studies have shown that the multivalent binding activity of Gal-1 and other galectins is able to cross-link carbohydrates and glycoconjugates [42,43]. For instance, Gal-1 cross-linking has the ability to massively redistribute a diverse population of glycoproteins on the cell surface of T cells and segregate them into membrane microdomains [44]. Gal-1 has also been associated with the assembly and remodelling of the extracellular matrix [45], and has been shown to bind to various components present there, especially those containing polylactosamine chains, such as laminins [45,46]. In this respect, *LAMC1* knockdown in NCH125 LGALS1 KO cells, or *LGALS1* knockdown in HeLa LAMC1 KD cells did not further decrease H-1PV cell uptake (Figure 6), suggesting that laminins and Gal-1 may cooperate in the early steps of H-1PV infection presumably by performing overlapping roles. However, as there is still residual internalisation of H-1PV (Figure 3C), it is likely that H-1PV may use alternative pathways to enter cells and that other unidentified cell factors are involved in this process. On the other hand, other laminins with or without the laminin-γ1 chain may contribute to residual H-1PV entry, independently from Gal-1. Future studies are needed to further characterize the pathways involved in H-1PV cell binding-entry and to determine whether other cellular factors besides laminins and Gal-1 are involved in these events.

Gal-1 and galectins in general have been described as playing important roles in different aspects of various viral infections, leading to their promotion or inhibition. For instance, Gal-1 stabilised the binding of HIV-1 to CD4+ T cells by cross-linking the viral gp120 and the host CD4 receptor, thereby helping HIV-1 to infect these cells [21]. Furthermore, soluble Gal-1 enhanced the uptake of HIV-1 by monocyte-derived macrophages, whereas Gal-3 had no effect on infection [47]. Enterovirus 71 is another example where Gal-1 has a supporting role. Gal-1 facilitates infection by interacting with the carbohydrate residues in VP1 and VP3 domains, leading to a more efficient release and dissemination of virus to other cells [48]. During Nipah virus (NiV) infection, Gal-1 enhances virus cell attachment to primary human endothelial cells [49]. However, later in the NiV replication cycle, Gal-1

seems to exert an inhibitory effect. Indeed, Gal-1 specifically binds to the viral glycoproteins NiV-F and NiV-G, which are responsible for cell-cell fusion and syncytia formation, thus blocking virus infection [50,51]. The inhibitory effect of Gal-1 is also observed in Influenza A infection, both in vitro and in vivo. Gal-1 binds directly to the envelope glycoproteins, stopping influenza from inducing hemagglutination and thereby impairing infectivity. Accordingly, influenza infection led to poorer survival rates in *LGALS1* KO mice than in wild-type mice [22].

Apart from their role in virus infections, galectins are also linked to apoptosis, angiogenesis, cell migration and tumour-immune escape [52]. In particular, high levels of Gal-1 are associated with cancer progression, poor prognosis and recurrence (reviewed in [34]). Several cancer types have been implicated, including gastric cancer [53], ovarian cancer [54], pancreatic cancer [55] and GBM [56]. In agreement with previous studies, our in silico analysis revealed that GBM presents significantly higher levels of *LGALS1* than normal tissues, and that *LGALS1* expression increases from grade II to IV gliomas (Figure S4). In terms of survival, high *LGALS1* expression is associated with a poor prognosis in glioma (Figure S5). To complement the bioinformatic analysis, we assessed a cohort of 122 patient biopsies by immunohistochemistry. We showed that Gal-1 protein levels vary across GBM with higher levels found in biopsies from patients with recurrent versus primary GBM, while in normal tissues the levels were relatively low (Figure 7). These findings corroborate previous studies showing that elevated levels of Gal-1 are associated with GBM progression [57–59]. They also further support the use of H-1PV to treat GBM, especially those cases with high Gal-1 protein content, given the key role that this protein has in virus entry and oncolysis.

We also found a correlation between *LGALS1* expression levels and the ability of H-1PV to induce oncolysis in 59 cancer cell lines (Figures 8 and 9). These results suggest that tumours with elevated *LGALS1* expression levels are likely to be more susceptible to H-1PV oncolytic activity. Building on these findings, we show that, while virus attachment is unaffected, virus entry is enhanced in the U251, LN308, U87 and A172-MG semi-permissive cell lines when H-1PV is administrated together with recombinant Gal-1 (Figure 9). Under these conditions (2 h from infection), H-1PV is still likely at the very early stages of infection, strongly supporting that Gal-1 plays a role in the H-1PV cell entry process. In agreement with enhanced infection, we found that susceptibility of these semi-permissive glioma cells to H-1PV oncolytic activity increases upon addition of exogenous Gal-1, suggesting that a certain level of Gal-1 is required for an efficient and productive H-1PV infection. Together, these findings support the idea that Gal-1 may represent a limiting factor for H-1PV oncolysis, and therefore, that tumours with high Gal-1 expression are more likely to respond to H-1PV treatment.

A similar correlation was previously shown for *LAMC1* where cell lines highly expressing this gene were found to be more susceptible to H-1PV oncotoxicity [14]. Anticorrelation analysis of *LAMC1* obtained in the previous work (Pearson correlation *R* = −0.52, *R*<sup>2</sup> = 0.27) stated a slightly higher anti-correlation than the one obtained in this re-analysis (*LAMC1 R* <sup>=</sup> −0.433, *<sup>R</sup>*<sup>2</sup> = 0.188). Using the current datasets, *LGALS1* anti-correlation (*LGALS1*, *R* = –0.510, *R*<sup>2</sup> = 0.260) is moderately higher than that of *LAMC1*. Update of the datasets with acquisition of novel mRNA sequencing data may account for these differences. Interestingly, the anti-correlation is strongest when the *LGALS1* and *LAMC1* expression levels are analysed together, suggesting that their combined expression analysis may better predict the success of H-1PV infection against a certain tumour. This is in line with the concept that both genes play an important role in H-1PV infectivity. Our results also present new scenarios for treatment in which exogenous administration of recombinant Gal-1 could constitute a promising adjunct in H-1PV-based therapies. However, given the role of Gal-1 in carcinogenesis [60], its possible use with H-1PV must be carefully evaluated.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/v14051018/s1, Figure S1: LGALS1 and LGALS3 knockdown in HeLa, NCH125 and BxPC3 cell lines. Figure S2: Cell proliferation of NCH125 Control versus NCH125 LGALS1 KO. Figure S3: Depletion of both *LAMC1* and *LGALS1* in NCH125 and HeLa cells. Figure S4: *LGALS1* overexpression in high-grade GBM. Figure S5: Higher levels of *LGALS1* are associated with poor prognosis in glioma. Figure S6: NCI60 database retrieved *LGALS1* and *LAMC1* expression levels showed anti-correlation with EC50 values and a slight increase in the ability to predict susceptibility to H-1PV induced oncotoxicity. Figure S7: CCLE database that retrieved *LGALS1* and *LAMC1* expression levels showed anti-correlation with EC50 values and showed a slight increase in the ability to predict susceptibility to H-1PV induced oncotoxicity.

**Author Contributions:** T.F. and A.K. designed, performed experiments and analysed the data; C.B. performed confocal microscopy analysis; J.A.H., L.A.R.Y. and H.M. provided and performed the GBM tissue microarray; R.R. and B.N. carried out the Nanostring analysis. P.V.N. performed correlation analysis. T.F., A.K. and A.M. wrote the manuscript and prepared figures. A.M. designed, secured funding, participated in data analysis, coordinated and supervised the research. All authors have read and agreed to the published version of the manuscript.

**Funding:** This study was supported initially by a seeding grant from Institut National du Cancer (INCA) and at later stages by a grant from ORYX GmbH to A.M. Our deepest gratitude also goes to André Welter, the Luxembourg Cancer Foundation and Télévie for supporting oncolytic virus immuno-therapy.

**Institutional Review Board Statement:** The tissue microarray prepared from GBM biopsies was approved by the Regional Ethics Committee of Bergen, (Norway) with the number 2017/2505.

**Informed Consent Statement:** Informed consent was obtained from all subjects involved in the study.

**Data Availability Statement:** All relevant data supporting the findings of this study are available within the paper and its Supplementary material. All other data are available from the corresponding author on request. Figure 8, Figure S6 and Figure S7 were generated employing the data sets publicly available at the CellMiner™ (https://discover.nci.nih.gov/cellminer, accessed on 23 January 2021) and at The Cancer Cell Line Encyclopedia portals (https://portals.broadinstitute.org/ccle, accessed on 23 January 2021). Figures S4 and S5 were generated using the Glio-Vis data portal (http://gliovis.bioinfo.cnio.es/, accessed on 23 January 2021).

**Acknowledgments:** We thank Barbara Leuchs (DKFZ, Heidelberg, Germany) for kindly providing the H-1PV capsid monoclonal antibody. We would also like to show our gratitude to Tiina Marttila for producing both the H-1PV wild-type and the recH-1PV-EGFP viruses and Anabel Grewenig for technical assistance in Western blot analysis. We thank Jean Rommelaere, Assia Angelova and Marcelo Ehrlich for fruitful discussion. We are also grateful to Caroline Hadley (INLEXIO) for critically reading the manuscript.

**Conflicts of Interest:** An international patent application protecting some of the results described in this article was submitted in November 2018 with T.F., A.K. and A.M. as co-inventors. A.M. is an inventor of several H-1PV-related patents/patent applications.

#### **References**


### *Article* **A Functional Minigenome of Parvovirus B19**

**Alessandro Reggiani, Andrea Avati †, Francesca Valenti ‡, Erika Fasano, Gloria Bua, Elisabetta Manaresi and Giorgio Gallinella \***

> Department of Pharmacy and Biotechnology, University of Bologna, 40138 Bologna, Italy; alessandro.reggiani5@unibo.it (A.R.); andrea.avati@student.unisi.it (A.A.); francesca.valenti@ior.it (F.V.); erika.fasano2@unibo.it (E.F.); gloria.bua2@unibo.it (G.B.); elisabetta.manaresi@unibo.it (E.M.)

**\*** Correspondence: giorgio.gallinella@unibo.it


**Abstract:** Parvovirus B19 (B19V) is a human pathogenic virus of clinical relevance, characterized by a selective tropism for erythroid progenitor cells in bone marrow. Relevant information on viral characteristics and lifecycle can be obtained from experiments involving engineered genetic systems in appropriate in vitro cellular models. Previously, a B19V genome of defined consensus sequence was designed, synthesized and cloned in a complete and functional form, able to replicate and produce infectious viral particles in a producer/amplifier cell system. Based on such a system, we have now designed and produced a derived B19V minigenome, reduced to a replicon unit. The genome terminal regions were maintained in a form able to sustain viral replication, while the internal region was clipped to include only the left-side genetic set, containing the coding sequence for the functional NS1 protein. Following transfection in UT7/EpoS1 cells, this minigenome still proved competent for replication, transcription and production of NS1 protein. Further, the B19V minigenome was able to complement B19-derived, NS1-defective genomes, restoring their ability to express viral capsid proteins. The B19V genome was thus engineered to yield a two-component system, with complementing functions, providing a valuable tool for studying viral expression and genetics, suitable to further engineering for purposes of translational research.

**Keywords:** parvovirus B19; synthetic genome; genetic engineering; replicon unit; functional complementation

#### **1. Introduction**

Within the family *Parvoviridae* [1], Parvovirus B19 (B19V) is a human pathogenic virus of clinical relevance, responsible for transient or persistent erythroid aplasia, infectious erythema, arthropathies, myocarditis and intrauterine infections, among others [2,3]. The variability in the pathogenetic processes and the resulting clinical outcomes of diverse nature and severity depend on a complex interplay between the viral properties, the characteristics of target cells in the different tissues, and the physiological status and immune response of infected individuals. B19V has a marked tropism for erythroid progenitor cells (EPCs) in the bone marrow, both susceptible and permissive depending on their differentiation and proliferation state [4]. In EPCs, infection normally induces cell cycle arrest and apoptosis [5], thus causing a temporary block in erythropoiesis which can become clinically relevant [6]. Different non-erythroid cell types, including endothelial, stromal, or synovial cells, are also susceptible but mainly non-permissive. In these cells, infection can trigger inflammatory responses and consequent tissue damage [7], and generally results in long-term persistence of viral DNA within tissues [8].

Investigation of viral genetics is fundamental to better understanding of the B19V replication cycle, the virus–cell interaction in different environments, the pathogenetic

**Citation:** Reggiani, A.; Avati, A.; Valenti, F.; Fasano, E.; Bua, G.; Manaresi, E.; Gallinella, G. A Functional Minigenome of Parvovirus B19. *Viruses* **2022**, *14*, 84. https://doi.org/10.3390/v14010084

Academic Editor: Luis Martinez-Sobrido

Received: 6 December 2021 Accepted: 29 December 2021 Published: 4 January 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

processes underlying the wide range of associated diseases, and the devising of more efficient antiviral strategies [9,10]. The B19V genome is composed of two inverted, terminal repeated regions (ITR) of imperfect palindromic sequence, 383 nt long, flanking a unique internal region (IR), 4830 nt long, containing all the coding sequences (Figure S1). The role of ITRs is crucial. The ability of palindromic sequences to fold in self-priming, hairpin secondary structures, and the presence of specific cis-recognition sequences acting as origins of replication, allow replication of the viral DNA through a rolling hairpin mechanism [11–13]. The activity of the unique transcription promoter (P6) at the left end of the internal region also depends critically on regulatory sequences within the upstream ITR [14–18]. Within the IR, distribution of splicing and cleavage-polyadenylation recognition sequences along the genome ensures coordinate processing of the pre-mRNA to a set of mature mRNAs [19–22]. Functionally, these can be divided into a set expressed from the left side of the genome, mainly coding for NS1 protein in an early phase of replication, and a set expressed from the right side of the genome, mainly coding for the structural VP proteins in the late phase of replication [23–25].

NS1 protein is the major non-structural protein, essential to virus replication and central for interaction with host cell components [9]. It is involved in the replication of the B19V genome, by its capacity to bind to specific recognition sites in the terminal regions, and by its endonuclease and helicase activities effecting ITR terminal resolution and strand unwinding. It is involved in viral transcription, enhancing activity of the P6 promoter by its trans-activating domains, thus promoting overall B19V genome expression. Besides, NS1 is a heterologous trans-activator of cellular genes, therefore inducing alterations in the cellular environment. It has a role in regulating progression through the cell cycle and in inducing apoptosis, therefore contributing to the pathogenesis of B19V infection. Given all this, NS1 is a matter of relevant interest in studying B19V, not least as a pharmacological target [26,27]. Viral capsid proteins VP1 and VP2 assemble to form a capsid shell of 22 nm in diameter, arranged in a T = 1 icosahedral structure [28]. Being translated from the same coding frame, both proteins share a common region that forms the core shell, while the VP1 protein, about 5% in abundance, possesses an additional N-terminal region, VP1u, crucial for cell recognition, attachment and penetration [29,30].

Information can be obtained from experiments involving engineered genetic systems in appropriate in vitro cellular models, since cloned forms of the B19V genome can be competent for replication and constitute an effective tool for studying the viral lifecycle and its interaction with target cells [31,32]. Previously, a model system was established by a novel synthetic strategy [33]. A reference genome of defined consensus sequence was designed, synthesized and cloned in a complete and functional form in a plasmid vector. Such a genome was able to replicate and produce infectious viral particles in a producer/amplifier cell system, the myeloblastoid UT7/EpoS1cells, allowing generation and further propagation of virus in EPC cell cultures. Replicative competence was linked to preservation of sequence integrity and asymmetry within the terminal regions, while preservation of the complete internal region ensured maintenance of the cis-acting signals required for regulation of viral genome expression. Therefore, the full proteome of B19V could be co-ordinately expressed and novel infectious viral particles produced. Further investigation is required to assess the flexibility of this system to genetic manipulation, and its potential as an advanced tool for basic research and translational applications.

With this aim, the main objective of the present experiments was to create a genetic unit derived from the complete, competent cloned B19V genome, simplified to a potential replicon unit. In this minigenome, the terminal regions were maintained in a form able to sustain viral replication, while the internal region was clipped to contain only the left-side genetic set. This would only allow the production of the subset of mRNAs corresponding to the early phase, including the mRNAs for NS1 protein. Following transfection in UT7/EpoS1 cells, replication, transcription and production of NS1 protein were monitored to assess the functional competence of the minigenome. Further, the capacity of this minigenome to complement defective forms of B19 genome rescuing production of viral capsid proteins and the possibility of producing transducing viral particles was also tested.

#### **2. Materials and Methods**

#### *2.1. Molecular Cloning*

Experiments were carried out on the previously established pCK10 and pCH10 plasmid clones containing the reference B19V EC sequence (GenBank KY940273) [33]. Plasmids pCK10 and pCH10 contain as inserts B19V EC segments, including the complete internal region and extension of both ITRs beyond the sites of dyad symmetry (pCK10, nt. 136–5461) or up to the sites of dyad symmetry (pCH10, nt. 184–5413).

The derived pCK10-pAs1 and pCH10-pAs1 plasmids were obtained by deletion of the genomic segment between nt. 2813–5169. To the purpose, a synthetic segment of appropriate sequence was transferred into pCK10 and pCH10 to replace the original insert by cloning using the XmaI and BssHII sites at positions 2251 and 5413. The derived pCH10-A1.1 and pCH10-A1.2 plasmids were obtained by deletion of the genomic segments between nt. 588–2089 and 588–2209, respectively. Synthetic segments of appropriate sequence were transferred into pCH10 to replace the original insert by cloning using the BssHII sites and BamHI sites at positions 184 and 4076.

Synthetic DNA inserts were obtained from Eurofins Genomics. Restriction endonuclease (RE) and ligase enzymes were obtained from Thermo Fisher Scientific and used according to manufacturer's directions. Plasmid clones were maintained in SURE bacterial cells (Agilent Technologies, Santa Clara, CA, USA) under ampicillin selection and growth in LB medium at 30 ◦C. Plasmid DNA purification was performed by PureYield Plasmid Midiprep (Promega, Madison, WI, USA). Inserts used for transfection assay were amplified by PCR by using the Expand High Fidelity System (Roche, Basel, Switzerland) as described, further purified by using Wizard SV Gel and PCR clean-up system (Promega) and quantified by UV absorbance determination.

#### *2.2. Cell Culture*

UT7/EpoS1 cells, obtained from KE Brown [34], were cultured in IMDM (Cambrex, East Rutherford, NJ, USA), 10% FCS and 2 U/mL Epo α (Eprex, Janssen, Beerse, Belgium), at <sup>37</sup> ◦C and 5% CO2. Cells were kept in culture at densities between <sup>2</sup> × 105–1 × 106 cells/mL and used for transfection experiments when at a density of 3 × <sup>10</sup><sup>5</sup> cells/mL. Erythroid progenitor cells (EPCs) were generated in vitro from peripheral blood mononuclear cells (PBMC) obtained from the leukocyte-enriched buffy coats of healthy blood donors, from the Immunohematology and Transfusion Service, S. Orsola-Malpighi University Hospital, Bologna (http://www.aosp.bo.it/content/immunoematologia-e-trasfusionale; authorization 0070755/1980/2014, issued by Head of Service). Availability was granted under conditions complying with Italian privacy law. Neither specific ethics committee approval nor written consent from donors was required for this research project. In vitro culture was carried out following established protocol [35], and cells were used for infection experiments at day 8 of in vitro growth and differentiation.

#### *2.3. Transfection and Infection*

UT7/EpoS1 cells were transfected by using the Amaxa Nucleofection System (Lonza, Basel, Switzerland), with V Nucleofector Reagent and T20 program setting, at a ratio of 1 μg insert DNA for 10<sup>6</sup> cells. Following transfection, the cells were incubated at 37 ◦C and 5% CO2 in complete medium at an initial density of 10<sup>6</sup> cells/mL, until collection at the indicated time points. Cells and cell-free supernatants were separated by centrifugation at 4000 rpm for 5 min in microfuge (Eppendorf, Hamburg, Germany), then fractions were used for analysis and/or successive infection experiments.

For infection experiments, cell-free supernatants obtained from transfected UT7/EpoS1 cells were added to EPCs cells at a ratio of 100 <sup>μ</sup>L for 1 × <sup>10</sup><sup>6</sup> cells. Infection was carried out at 37 ◦C for 2 h, then cells were washed free of inoculum and expanded in complete medium at 37 ◦C and 5% CO2 at an initial density of 106 cells/mL, until collection at the indicated time points and subsequent processing as described.

#### *2.4. Quantitative Molecular Analysis*

Experimental samples were processed for total nucleic acid purification by using the Viral Total Nucleic Acid kit for the Maxwell 16 extractor (Promega), then quantitative determination of B19V nucleic acids (viral DNA, total mRNA, mRNA subsets) was carried out by qPCR and qRT-PCR according to previously established protocols [23,24]. Genomic DNA coding for 18S rRNA (rDNA) was amplified for calibration with respect to cell copy number. Absolute quantification of both viral DNA and total viral RNA was obtained by using the primer pair R2210–R2355, located in the central exon of B19V genome, while determination of the relative abundance of the different subsets of viral transcripts was obtained by using a selected array of primer pairs, as indicated in Table 1. For the DpnI Assay, DNA previously treated with either EcoRI or EcoRI+DpnI restriction enzymes was amplified by using primers encompassing DpnI site at position 1801 on the B19V genome, and the fraction of DNA not cleaved by the enzyme determined by qPCR analysis and absolute quantitation with respect to an external calibration curve.

**Table 1.** Primer combinations used in the qPCR and qRT-PCR assays for the detection and quantitative evaluation of B19V nucleic acids. See also Figure S1 for primer location on B19V genome.


#### *2.5. IIF and Cytofluorimetric Analysis*

For detection of viral proteins by immunofluorescence, aliquots of 5 × <sup>10</sup><sup>4</sup> cells were spotted on glass slides and fixed with 1:1 acetone:methanol for 10 min at −20 ◦C. For detection of NS protein, cells were incubated with the human monoclonal antibody MAb1424 (kindly supplied by Susanne Modrow) (1:100 in PBS/FCS 10%), then with an antihuman FITC-conjugated secondary antibody (Dako, 1:20 in PBS/FCS 10%). For detection of VP proteins, cells were incubated with a monoclonal mouse antibody against VP1 and VP2 proteins (MAb8293, Chemicon, Merck Millipore, Milan, Italy) (1:200 in PBS/BSA 1%), then with AlexaFluor488 anti-mouse secondary antibodies (Life Technologies, Monza, Italy) (1:1000 in PBS/BSA 1%). Cell populations were also analysed for expression of viral proteins by using flow cytometry (FACSCalibur, Becton Dickinson, Milan, Italy). Aliquots of 106 cells were fixed in PBS/formaldehyde 0.5% O/N at 4 ◦C, permeabilized in PBS/saponin 0.2% at RT while rocking for 45 min and incubated in suspension with antibodies diluted in PBS/FCS 2% (1:100 NS primary; 1:40 anti-human FITC secondary). Data were analysed using the Cell Quest Pro Software (Becton Dickinson).

#### **3. Results**

#### *3.1. Design and Construction of a B19V Minigenome*

In the design of a B19V minigenome with potential replicative activity, the rational requirements were: (i) to preserve both terminal regions up to the sites of dyad symmetry, retaining the capacity of hairpin formation; and (ii) to preserve the internal region extending up to the pAp1 proximal cleavage-polyadenylation signal, as a gene cassette with potential for coding for the NS1 protein, while eliminating the genomic region coding for the viral capsid proteins. To this end, a large deletion was operated in the right-side of genome,

and a novel chimeric cleavage-polyadenylation signal created, named pAs1, joining the upstream cis-elements of pAp1 and the downstream cis-elements of pAd (Figure 1).

**Figure 1.** (**A**) Map of B19V genome. ITR: inverted terminal repeats (-, site of dyad symmetry). IR: internal region and relevant cis-acting functional sites (P6, promoter; pAp1, pAp2, proximal cleavagepolyadenylation sites; pAd, distal cleavage-polyadenylation site; D1, D2, splice donor sites; A1.1, A1.2, A2.1, A2.2, splice acceptor sites). Coding sequences for viral NS, VP and smaller non-structural proteins are aligned to map. **Δ**: deletion to create a novel cleavage-polyadenylation signal (pAs1). (**B**) Map of B19V derived minigenome; simplified transcription map, indicating the two classes of mRNAs (mRNA 1–2), with alternative splicing forms (dashed lines) and related coding potential. (**C**) Sequence at the novel pAs1 cleavage-polyadenylation site (USE—pAp1, upstream element to pAp1 [19]).

For the construction of the minigenome, a synthetic gene segment encompassing the designed deletion substituted by the novel pAs1 sequence was synthesised and inserted in order to replace the original sequence in the previously established pCK10 and pCH10 plasmids [33]. The viral insert in pCK10 preserves both terminal regions extending beyond the site of dyad symmetry; however, attempts at cloning in pCK10 only yielded unstable plasmid clones, with deletion of the palindromic sequence in the right-hand terminal region. The viral insert in pCH10 preserves both terminal regions extending up to the site of dyad symmetry; in this case, a stable plasmid clone was successfully obtained, named pCH10-pAs1.

#### *3.2. Functional Competence of the B19V Minigenome*

From the pCH10 plasmid, three genomic inserts of different extension can be obtained, differing in their functional competence: CH10, corresponding to the whole cloned insert, extending in the terminal regions to the sites of dyad symmetry (nt. 184–5413); CI0, extending in the terminal regions within the sites of dyad symmetry (nt. 245–5474); and CJ0, extending in the terminal regions only to the start of palindromes (nt. 366–5231). From the pCH10-pAs1 plasmid, three genomic inserts of corresponding extension could also be obtained by analogy: CH10-pAs1, CI0-pAs1, and CJ0-pAs1. The biological activity and functional competence of pCH10 and pCH10-pAs1 derived inserts was comparatively analysed following transfection in UT7/EpoS1 cells. Genomic inserts were obtained by means of in vitro amplification, then purified inserts were used to transfect UT7/EpoS1 cells. At 8- and 24-h post-transfection (hpt), aliquots of cell culture were sampled for quantification of viral nucleic acids (DNA, mRNAs) by qPCR and qRT-PCR (Figure 2, Table S1A), and detection of the NS protein by IIF and cytofluorimetric analysis.

**Figure 2.** Viral nucleic acids in UT7/EpoS1 cells, transfected with CH10 and CH10-pAs1 derived inserts. Log amounts of target copies (viral DNA, total RNA, NS1 mRNA), normalized to 105 cells, at 8 and 24 hpt. Mean and std of duplicate determinations for two different experiments. Two-way ANOVA, Bonferroni post-test: \*\*\*, *p* < 0.001; \*, *p* < 0.05.

The amount of DNA, either at 8 or 24 hpt, was comparable for all tested inserts, indicating a similar transfection efficiency. Due to the large quantity of input DNA used in transfection, no significant temporal variation in DNA amount was observed for any of the tested inserts, apart from a general decrease from 8 to 24 hpt (mean 0.70 Log, range −1.1–0.57), likely to be due to progressive degradation of exogenous DNA. De novo synthesis of transfected DNA was assessed in a parallel experiment, by transfection of inserts directly excised from plasmids and tested for *Dam* methylation pattern and resistance to DpnI cleavage, both by a qPCR assay and a Southern Blot analysis. In this way, it was possible to investigate CH10 and CH10-pAs1, excised using BssHII, CI0 and CI0-pAs1, excised using AccIII, but not CJ0 and CJ0-pAs1, because of lack of corresponding RE sites.

By qPCR, DNA excised from plasmids was resistant to 1.7% and 1.4% for CH10 and CH10-pAs1, and 1.6% and 1.3% for CI0 and CI0-pAs1. DNA obtained from transfected cells at 24 hpt was resistant to 11.7% and 15.8%, and 8.8% and 19.0%, respectively. By Southern Blot (Figure 3), at 24 hpt, bands corresponding to full-length, DpnI resistant DNA were also observed for all transfected inserts. Data thus obtained are consistent with the maintenance of the replicative competence of transfected inserts, at least for CH10-and CI0-derived inserts, both the complete genomes and the derived minigenomes, a property likely to be due to the preservation of sequence symmetry within the terminal regions and implying a hairpin-independent priming of DNA synthesis.

All transfected inserts showed a sustained transcriptional activity. Viral mRNAs were detected for all inserts at both time-points post-transfection, with a general increase from 8 to 24 hpt of total mRNA (mean 0.56 Log, range −0.01–1.33), and to a lesser extent of NS mRNA (mean 0.08 Log, range −0.36–0.51), significant for CH10-pAs1 only. Although with some variability, overall results attested the early onset and maintenance of viral transcription, implying processing of pre-mRNA at the novel chimeric pAs1 cleavagepolyadenylation site, and preservation of a balanced usage of splicing signals. In fact, a typical ratio of mRNAs pertaining to the left-side genome was produced, including both the unspliced mRNAs coding for NS protein (mRNA 1 in Figure 1), approximately 1% of

total viral mRNAs, and the more abundant spliced mRNAs (mRNA 2 in Figure 1), in a pattern analogous to that observed in the early phase of the replicative cycle.

**Figure 3.** Southern Blot Analysis of B19V DNA obtained from UT7/EpoS1 cells transfected with inserts CH10, CI0 and CH10-pAs1, CI0-pAs1, collected at 24 hpt. Samples were treated by RE DpnI to distinguish de novo synthesized viral DNA (\*) based on different *dam* methylation pattern and sensitivity to RE DpnI. MwM: molecular weight marker III, Dig-labelled (Roche). Southern Blotting and hybridization using a full-length digoxigenin-labelled DNA probe was carried out as described [33].

The expression of NS protein was monitored by IIF and cytofluorimetric analysis, sampling transfected cell cultures at 8 and 24 hpt. By IIF (Figure 4), NS protein was already observed at 8 hpt, and at 24 hpt the number of positive cells and signal intensity both increased. Distribution of the protein within the cells showed the same nuclear/cytoplasmic pattern both for all complete or derived minigenome inserts.

By cytofluorimetric analysis (Figure 5), a quantitative assessment of cells expressing NS1 protein was obtained at 24 hpt. The percentage of positive cells was in the range 0.2–1.3% for the complete inserts and increased to 2.6–5.0% for the derived minigenomes, the highest increase was observed for the CI0/CI0-pAs1 combination. Altogether, experiments suggest that the lower genetic complexity of modified genomes promoted progressive expression and accumulation of NS protein in an increasing fraction of the cell population.

**Figure 4.** UT7/EpoS1 cells transfected with the indicated inserts were sampled at 24 hpt, and NS1 protein was detected by IIF. Original magnification 400×.

**Figure 5.** UT7/EpoS1 cells transfected with the indicated inserts (Ctrl, no DNA control) were sampled at 24 hpt, and cell population was analysed by cytofluorimeter to determine the percentage of NS1 expressing cells. Dot plot graph on gated cell population for FSc and NS1 FITC. Reported percentage values of positive cells and geometric mean fluorescence intensity (MFI) for positive subpopulations reported as the average result of two independent determinations.

#### *3.3. Functional Complementation of the B19V Minigenome*

The capacity of the CH10-pAs1 minigenome to provide complementing functions through expression of NS1 protein was thereafter tested in a subsequent series of experiments. To the purpose, a set of modified B19V genome clones was designed, defective for the coding sequence of NS protein. In particular, the pCH10 clone was modified by deletion, removing the genomic region corresponding to the first intron within the NS gene, from the splice donor site D1 to the two possible alternative splice acceptor sites A1.1 and A1.2. The obtained plasmids, named CH10-A1.1 and CH10-A1.2, retained the cis-acting sequences directing alternative cleavage-polyadenylation at pAp and pAd sites, sequences regulating alternative splicing of the distal introns at D2 and A2.1/2 sites, and all of the coding sequences for the VP1, VP2 and 11 kDa proteins (Figure 6).

**Figure 6.** (**A**) Map of B19V genome, see Figure 1. **Δ**: deletion to remove first intron (A1.1/2). (**B**) Map of B19V derived minigenomes; simplified transcription map, indicating the four classes of mRNAs (mRNA 2–5), with alternative splicing/cleavage forms (dashed lines) and related coding potential.

To evaluate the functional competence and possible complementation effects for these defective genomes, inserts obtained by means of in vitro amplification were transfected in UT7/EpoS1 cells, alone or in co-transfection with CH10-pAs1 as helper plasmid. At 24 hpt, aliquots of cell culture were sampled for quantification of viral nucleic acids (DNA, mRNAs) by qPCR and qRT-PCR (Figure 7, Table S1B), and detection of the NS1 and VP proteins by IIF.

No significant differences were observed in the amounts of viral DNA, but relevant information was obtained by quantification of viral RNA. By comparison to the reference CH10 insert, insert CH10-pAs1 confirmed its high transcriptional activity and correct mRNA processing, with an abundance of about 1% unspliced mRNAs coding for NS protein. The NS defective, CH10-A1.1 and -A.2 inserts showed a reduced (−3 Log) basal transcriptional activity, which allowed detection of only the proximally cleaved mRNAs (mRNA 2 in Figure 6), and not of any distally cleaved mRNA (mRNAs 3–5 in Figure 5). Cotransfection of these inserts with CH10-pAs1 inset led to functional complementation, restoring expression from the NS-defective genomes, as shown by the detection of A1.1 and A.2 derived mRNAs, to amounts only about 1 Log lower than what observed for the reference CH10 insert. By composition, the most abundant mRNAs were still the proximally cleaved mRNA species (mRNA 2), which were contributed by both CH10-pAs1 (following splicing) and A1.1/2 (following cleavage at pAp); abundance of NS mRNA (mRNA 1), contributed only by CH10-pAs1, pAd cleaved mRNAs (mRNA 3–5) and VP mRNAs (mRNA 3–4), contributed only by A1.1/2, were in all cases in the order of 1% of total mRNAs.

**Figure 7.** Viral nucleic acids in UT7/EpoS1 cells, transfected/co-transfected with CH10, CH10-pAs1 and CH10-A1.1/2 inserts. Log amounts of target copies (viral DNA, total RNA, NS1 mRNA, pAd cleaved RNA, VP RNA), normalized to 10<sup>5</sup> cells, at 24 hpt. Mean and std of duplicate determinations for two different experiments.

By IIF analysis (Figure 8), expression of both NS and to lesser extents VP proteins was confirmed for the CH10 insert. The expression of NS protein was also confirmed for CH10-pAs1, and not detected in the case of CH10-A1.1/2, as expected. Expression of VP proteins from CH10-A1.1/2 inserts alone was not observed. Cotransfection of CH10 pAs1 and A1.1/2 preserved expression of NS protein from CH10-pAs1 and restored the expression of VP proteins from CH10-A1.1/2, although for the latter detection was limited to a small number of cells. Altogether, data confirm that the NS protein produced by the CH10-pAs1 minigenome is functional in complementation of defective B19V genomes, restoring expression of the late set of mRNAs to a pattern similar to the standard expression profile of B19V genome and allowing production of capsid proteins.

**Figure 8.** UT7/EpoS1 cells transfected with the indicated inserts were sampled at 24 hpt, and NS1 or VP1/2 proteins were detected by IIF. Results obtained for insert CH10-A1.1 were analogous to what obtained for CH10-A1.2 (not in figure). Original magnification 400×.

#### *3.4. Extracellular Vehiculation of Minigenomes*

Following transfection of UT7/EpoS1 cells, measurable amounts of viral DNA were detectable in the cell culture medium until 6 days post-transfection, not associated to cells but partially resistant to nuclease treatment. The possibility that this genetic material could be transferrable to susceptible EPCs was investigated. For the purpose, CH10-pAs1 and CH10-A1.1/2 inserts were transfected in UT7/EpoS1 cells in the different combinations as described, and expression of NS and VP proteins first confirmed for all competent combinations. Then, after a 6 day course, the supernatant of transfected cell cultures was collected and added to in vitro differentiated EPCs cells, as a test system. After a further 48 h course of incubation, EPCs were collected and analysed for any presence of B19V DNA, RNA or expression of NS protein. For all experimental samples, a low measurable amount of DNA was found associated to EPCs (<102 copies/105 cells). However, no transcriptional activity could be detected in EPCs in any tested combination and no NS protein production could be observed. The obtained results imply that the vehiculation of genetic material from cells transfected with the CH10 derived clones may not be due to the formation of transducing viral particles, and that the process is not functionally relevant to a measurable extent. The hypothesis that such transfer should be attributed to simple carry-over in a nuclease-resistance form, or to the formation of extracellular vesicles or exosomes, and whether a limited number of transducing viral particles is actually produced, requires further investigation.

#### **4. Discussion**

In our present work, we designed and produced a B19V minigenome, derived from the complete, competent cloned B19V genome, simplified to a replicon unit. Following transfection in the UT7/EpoS1 cells, this minigenome still proved competent for replication, transcription and production of NS protein. Further, the B19V minigenome was able to complement B19-derived, NS-defective genomes, restoring their ability to express capsid proteins. The unique B19V genome was thus engineered to yield a two-component system, an element expressing the functional NS1 protein, the other the structural capsid proteins, with complementing functions.

In all engineered genetic elements, the terminal regions have been preserved up to the site of dyad symmetry and in opposite flip/flop configurations, thus meeting requirements for maintenance of replicative competence [12,33]. To this respect, interesting information has been obtained by comparing the activity of genomic inserts of different extension within the ITRs. De novo synthesis of transfected DNA could be demonstrated, as previously [33], for the complete CH10 and CI0 inserts, and this property was also conserved for the derived -pAs1 inserts. Hairpin-independent priming of DNA synthesis had been documented in a different experimental setting [11], and it is likely involved here.

The unique P6 promoter is present in the same position and pattern in all engineered clones, whereas the level of transcription of each one depends on the actual expression of NS protein, due to its strong trans-activating activity on its own promoter [14]. Transcription from the complete CH10, CI0, CJ0, and the derived -pAs1 inserts is detected at high levels already at an early time point (8 hpt), further increasing at a late time point (24 hpt). Transcription levels from the derived -pAs1 inserts is higher than the respective complete clones, but effects depending on the ITR extension are relatively minor. Conversely, sustained transcription from CH10-A.1/2 clones is detectable only upon complementation, and at the late time point.

CH10-pAs1 elements are simplified with respect to pre-mRNA processing. Reduction in the size and complexity of the transcriptional template and the introduction of a novel single cleavage-polyadenylation site at the end of the genome abrogates the early-late transcriptional switch typical of B19V, thus leading to sole accumulation of left-side cassette mRNAs [19,20]. Within these processes, retention of unaltered splicing signals ensures correct processing of pre-mRNAs and unaltered balance of unspliced, NS encoding to spliced mRNAs [25]. Thus, the net effect compared to that observed for complete clones is a sustained accumulation of NS-coding transcripts and overexpression of NS protein in a larger proportion of transfected cells.

Modifications introduced to obtain the defective CH10-A1.1/2 elements have different consequences. Reduction of the genetic template with deletion of the large left-side intron results in the absence of any NS coding mRNA, but both original cleavage-polyadenylation sites are maintained, therefore preserving a possible early-late switch in expression pattern. Moreover, the splicing signals maintained in the right gene cassette still allow alternative splicing events leading to the production of VP1, VP2 and 11 kDa encoding mRNAs [21,22]. In fact, experiments confirmed that such an mRNA set is produced when in the presence of complementing NS protein. However, compared to the complete genome, a relatively higher proportion of short (proximally cleaved, spliced) mRNAs is produced, likely because of the contribution from both transcriptional templates and prevalent pre-mRNA processing. As a consequence, the amount of VP-encoding mRNA obtained from CH10- A.1/2 templates is reduced to suboptimal levels, although enough to achieve production of VP proteins.

The present work reached relevant goals, while showing some critical limitations inherent to the system. A minigenome with characteristics of a replicon has been constructed, able to replicate and overexpress NS protein. Such minigenome may constitute a valuable tool for studying the function of NS protein within a simplified viral genetic contest, mainly its impact on cell functionality, or in the search of molecules with antiviral activity. A functional complementation between defective genomes was obtained, since the minigenome restored the ability of separate genetic units to express capsid proteins. This property opens the possibility of engineering the B19V genome with less stringent structural constraints, allowing for example to modify cis-acting genomic elements, mutagenize or add expression tags to the sequences coding for proteins, or insert heterologous reporter genes, to the purpose of a deeper characterization of B19V replication, expression and interaction in the cellular environment.

Ideally, the compresence in a same cell of two genetic units with complementing functions opens a possibility for packaging and generation of transducing viral particles. However, generation of transducing virus was not shown in our experiments, thus constituting the major limit of the present work. All transfection experiments have been conducted in the UT7/EpoS1 cell line, which is appropriate for research on B19V [36], but at the expense of a very low transfection efficiency. When transfected with a complete genomic insert, de novo produced virus can be subsequently amplified in primary EPCs to yield infectious virus at high titre. In the case of cotransfection of separate complementing units, transducing viral particles would not benefit of any possible subsequent amplification passage. While generation of engineered virus specifically targeting a selected cell population such as EPCs would be of translational interest, further intense research is required to achieve such goal.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10. 3390/v14010084/s1, Figure S1: B19V genome organization, transcription map and primer location. Table S1: Quantitation of viral nucleic acids.

**Author Contributions:** Conceptualization, A.R., G.B., E.M. and G.G.; Data curation, A.R., G.B. and E.M.; Funding acquisition, G.G.; Investigation, A.R., A.A., F.V. and E.F.; Methodology, A.R., A.A., F.V. and E.F.; Supervision, G.B., E.M. and G.G.; Writing—original draft, A.R. and G.G.; Writing—review & editing, G.G. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Italian Ministry of University and Research, grant PRIN 2017 9JHAMZ\_007 to G.G.

**Institutional Review Board Statement:** Leukocyte-enriched buffy coats of healthy blood donors, were obtained from the Immunohematology and Transfusion Service, S. Orsola-Malpighi University Hospital, Bologna (http://www.aosp.bo.it/content/immunoematologia-e-trasfusionale; authorization 0070755/1980/2014, issued by Head of Service). Availability was granted under conditions complying with Italian privacy law. Neither specific ethics committee approval nor written consent from donors was required for this research project.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

#### **References**

