Transbody against virus core protein potently inhibits hepadnavirus replication in vivo: evidence from a duck model of hepatitis B virus

Abstract

Background and Purpose

The therapeutic management of hepatitis B virus (HBV) infections remains challenging, and novel antiviral strategies are urgently required. The HBV transbody, a monoclonal antibody (MAb) against human HBcAg coupled with the trans‐activator of transcription protein transduction domain (TAT PTD), was previously shown to possess cell‐penetrating ability and potent antiviral activity in vitro. The purpose of the present study was to evaluate the antiviral activity of the HBcMAb‐TAT PTD conjugate in vivo in a duck model of HBV.

Experimental Approach

Female Peking ducks were injected i.p. with 0.03‐0.3 mg·kg⁻¹·day⁻¹ of the DHBV transbody (DHBcMAb‐TAT PTD conjugate) for 30 days. Serum DHBV DNA levels and liver DHBV DNA and covalently closed circular DNA (cccDNA) loads were determined at scheduled time points. Immunohistological examination of DHBcAg in the duck liver was also performed.

Key Results

The DHBV transbody significantly reduced the serum and liver DHBV DNA levels at doses of 0.1 and 0.3 mg·kg⁻¹·day⁻¹ and liver cccDNA levels at a dose of 0.3 mg·kg⁻¹·day⁻¹ after 30 days of treatment. The suppressive effects of the DHBV transbody at 0.3 mg·kg⁻¹·day⁻¹ on the serum and liver DHBV DNA and liver cccDNA levels remained significant, even at 15 days after treatment cessation. Similarly, immunohistochemistry of liver sections revealed decreased DHBcAg levels within hepatocytes 15 days after treatment termination.

Conclusions and Implications

The DHBV transbody inhibits DHBV replication and possesses potent anti‐DHBV activities in vivo. The cell‐permeable antibody against the virus core antigen may be developed as a novel treatment for patients with hepadnavirus infections.

Abbreviations

ALT

alanine aminotransferase

AST

aspartate aminotransferase

BUN

urea nitrogen

CRE

creatinine

DHBcMAb

DHBV core antigen monoclonal antibodies

DHBV

duck hepatitis B virus

HBcAg

hepatitis B virus core antigen

HBeAg

hepatitis B virus e antigen

HBV

hepatitis B virus

HPF

high‐power field

mAb

monoclonal antibody

MD

mean density

pgRNA

pregenome RNA

RBC

erythrocyte

scFv

single‐chain antibody

TAT PTD

trans‐activator of transcription protein transduction domain

VHH

variable domain of heavy chain of heavy‐chain antibody

WBC

leukocyte

Introduction

More than 350 million people are chronically infected with hepatitis B virus (HBV) worldwide, and this infection is responsible for 1 million deaths each year (Te and Jensen, 2010). Typically, chronic hepatitis and its complications, liver cirrhosis and hepatocellular carcinoma, are the causes of death in HBV infection. Several antiviral drugs have been approved for the treatment of HBV, including IFN‐α and nucleotide/nucleoside analogues. However, significant unresolved issues remain with the current drugs, such as moderate efficacy, dose‐dependent side effects and drug resistance (Rajbhandari and Chung, 2016). Therefore, the development of a safer and more potent therapeutic regimen is urgently needed for patients with chronic HBV infection.

During the HBV life cycle, the hepatitis B core antigen (HBcAg) is essential not only for nucleocapsid assembly but also for the regulation of viral replication. Several studies have shown that HBcAg is a good candidate for inhibition of HBV replication. Efficient viral inhibition by targeting HBcAg RNA or HBcAg was previously demonstrated with RNA interference (Ying et al., 2003), antisense oligonucleotides (Deng et al., 2015) and ribozymes (Feng et al., 2011), aptamers (Zhang et al., 2009), sulfamoylbenzamide derivatives (Campagna et al., 2013) and intrabodies, including single‐chain variable fragments (scFvs) and single‐domains [e.g. variable domain of heavy chain of heavy‐chain antibody (VHH)] (Yamamoto et al., 1999; Serruys et al., 2010).

Clinically, most hepatitis B patients produce an anti‐HBcAg antibody of high titre, which is only used as a diagnostic marker for HBV infection (Sherlock and Dooley, 1997). The anti‐HBcAg antibody does not show any antiviral effect because of its inability to enter virus‐infected cells. In our previous study, monoclonal antibodies (MAbs) against the core antigen of human HBV were coupled with the trans‐activator of transcription protein transduction domain (TAT PTD) to form transbodies, and we found that the HBV transbody decreased not only extracellular and intracellular HBV surface antigen, HBeAg and HBV DNA but also intracellular HBcAg, thereby effectively inhibiting HBV replication in HepG2.2.15 cells (Wang et al., 2015).

In this study, duck HBV (DHBV), another member of the hepadnaviridae family, which is closely related to human HBV, was used as an animal model for HBV (Schultz et al., 2004). DHBV core antigen MAbs (DHBcMAbs) were prepared and developed into a cell‐penetrable format, the DHBV transbody, by coupling the TAT PTD (residues 47–57:YGRKKRRQRRR) (Joliot and Prochiantz, 2004; Murriel and Dowdy, 2006) to DHBcMAb. The DHBV transbody was examined for its ability to inhibit DHBV in vivo in DHBV‐infected ducks.

Methods

Preparation of mouse DHBcAg MAb‐TAT PTD

A standard prokaryotic expression system with Escherichia coli BL21 as host strains and pET28a(+) (Invitrogen, Carlsbad, CA, USA) as the basic plasmid was used for the expression of the target protein DHBcAg. The DNA fragment encoding DHBcAg was amplified by PCR from pBR322/2DHBV (kindly provided by Dr Mason, Fox Chase Cancer Center, Philadelphia, PA, USA) and inserted into the BamHI/XhoI sites of pET28a(+) to yield the plasmid pET28a(+)/DHBV core. The recombinant plasmids were transiently transformed into E. coli BL21 bacteria, and expression was induced with isopropyl β‐D‐thiogalactopyranoside. The recombinant proteins were purified using Sepharose 4 Fast Flow and DEAE‐Sepharose Fast Flow (GE healthcare, Buckinghamshire, UK). The purified DHBcAg was identified using SDS‐PAGE with Coomassie blue staining (Figure 1A).

Figure 1.

SDS‐PAGE analysis of DHBcAg, DHBcMAb and the evaluation of DHBcMAb‐TAT PTD conjugate activity of DHBcAg binding. (A) SDS‐PAGE analysis of DHBcAg produced from recombinant plasmids pET28a(+)/DHBV core and E. coli BL21 for expression. Lane 1: purified DHBcAg showing a band of 30KDa; lane 2: precipitation of the expression of recombinant plasmids pET28a(+)/DHBV core; lane 3: supernatant of the expression of recombinant plasmids pET28a(+)/DHBV core; lane 4: precipitation of the expression of recombinant plasmids pET28a(+); lane 5: supernatant of the expression of recombinant plasmids pET28a(+); lane M: protein molecular weight marker. (B) SDS‐PAGE analysis of DHBcMAb using hybridoma technology. Lane 1: DHBcMAb before purification; lane 2: purified DHBcMAb; lane M: protein molecular weight marker. (C) Evaluation of DHBcMAb‐TAT PTD conjugate activity of DHBcAg binding. Binding of the DHBcMAb‐TAT PTD conjugate to DHBcAg by an ELISA using DHBcAg precoated on the plate and mouse anti‐TAT‐HRP as the detection reagent (n = 3).

Female BALB/c mice (8 weeks of age) were injected i.p., firstly with 50 μg of recombinant DHBcAg in complete Freund's adjuvant, then with 25 μg of DHBcAg in incomplete Freund's adjuvant 2 and 3 weeks later, and finally with 25 μg of DHBcAg alone 4 weeks later. Four days prior to hybridization, the mice were injected i.p. with the same dose of antigen alone. Myeloma SP2/0 cells were fused with splenocytes using the polyethylene glycol method. At day 15, cells producing anti‐DHBcAg were detected by enzyme‐linked immunosorbent assay (ELISA). Positive wells were cloned by the limiting dilution method.

Female BALB/c mice (8 weeks of age) were injected i.p., firstly with 0.5 mL of sterilized liquid paraffin and then with 0.5 mL of hybridoma cell suspension (1 × 10⁷ cells mL⁻¹) 1 week later. At day 10, ascites was collected. DHBcAg MAbs were purified from the ascites using n‐caprylic acid‐(NH₄)₂SO₄ precipitation. The precipitate was dissolved and then dialysed against PBS. Antibodies were purified by immunoaffinity chromatography using solid‐phase bound protein A (Figure 1B).

The purified DHBcAg MAbs were concentrated with polyethylene glycol 20 000 to one third of the original volume, and the protein concentration was quantified using the Coomassie brilliant blue G‐250 colorimetric method. The protein concentration was approximately 4 mg·mL⁻¹.

The preparation of DHBcMAb coupled with TAT PTD (Beijing Scilight Biotechnology Ltd. Co., Beijing, China) was performed according to the manufacturer's instructions using the Imject Immunogen EDC Conjugation Kits (Pierce, Holmdel, NJ, USA). Briefly, DHBcMAb was dissolved in 0.2 mL of deionized water, and equimolar TAT PTD with antibody was dissolved in 0.5 mL of conjugation buffer and added to 0.2 mL of antibody solution. Then, a threefold molar excess of 1‐ethyl‐3‐(3‐dimethylaminopropyl)carbodiimide hydrochloride (EDC) was added to the antibody/peptide solution, and the mixture was incubated for 2 h at room temperature. The DHBcMAb‐TAT PTD conjugate was purified by gel filtration using the column provided with the kits and eluted with 0.9 M NaCl.

DHBcMAb titres after purification, concentration and coupling with TAT PTD were detected using ELISA.

DHBcMAb‐TAT PTD conjugate‐binding ELISA

Microplates precoated with 0.1 mL of 1 μg·mL⁻¹ DHBcMAb were blocked with 0.2% BSA/PBS. After three washes with 0.1% Tween‐20/PBS, 0.1 mL of serially diluted DHBcMAb‐TAT PTD conjugates were added. Additionally, 0.1 mL of 10 μg·mL⁻¹ DHbcMAb, TAT PTD and PBS were added as controls. The plates were then incubated for 1 h at 37°C. Subsequently, the plates were washed three times with 0.1% Tween‐20/PBS, and incubated with 0.1 mL of serially diluted mouse anti‐TAT‐HRP conjugate (Xi'an Hua Guang Biology engineering Co., Xi'an, China) for 1 h at 37°C. HRP activity was determined using tetramethylbenzidine substrate. After the addition of 0.05 mL of 2.0 M H₂SO₄, the optical density was read at 450 nm.

Animals

Female Peking ducks (Anas platyrhynchos) 12 months old and weighing 0.9–1.1 kg were purchased from Duli Duck Farm of Shaanxi Province. Ducks were held at the animal house facilities in Xi'an Jiaotong University Health Science Centre and were maintained under normal daylight and fed with a standard commercial diet and water ad libitum. Blood samples were collected from the axillary vein. For liver biopsies, the ducks were anaesthetized with pentobarbital sodium (i.p., 30.0 mg·kg⁻¹) and laparotomized to obtain a small piece of the liver tissue (approximately 1 × 2 × 3 cm³ per duck). After closing the abdominal cavity, the breathing and heartbeat of the ducks were closely observed until completely awake. At the end of the experiment, the ducks were killed by decapitation by an experienced investigator. All animal experiments were carried out in accordance with the Guide for the Care and Use of Laboratory Animals (Institute of Laboratory Animal Resources). This project was approved by the animal care committee of School of Medicine, Xi'an Jiaotong University (protocol identification number No. XJTULAC2012‐226). Animal studies are reported in compliance with the ARRIVE guidelines (Kilkenny et al., 2010; McGrath and Lilley, 2015).

In vivo assays of the anti‐DHBV activity of DHBcMAb‐TAT PTD conjugate in ducks

After detection of DHBV DNA in blood samples, ducks with DHBV DNA > 1 × 10⁸ copies mL⁻¹ were randomized into seven groups (n = 6 per group). The liver tissue (1 × 2 × 3 cm³) of each duck in all groups was obtained by surgery and divided into two parts. One part was used for immunohistological examination of DHBcAg, and the other part was used for the determination of liver DHBV DNA and covalently closed circular DNA (cccDNA). The detected values were used as the baseline treatment parameters. After recovery for 10 days, three groups of ducks were injected i.p. with DHBcMAb‐TAT PTD conjugate at 0.03, 0.1 and 0.3 mg·kg⁻¹·day⁻¹ for 30 days. One group of ducks was treated p.o. with 10 mg·kg⁻¹·day⁻¹ of adefovir dipivoxil (GlaxoSmithKline, Middlesex, UK) for 30 days as a positive control, and one group of ducks was injected i.p. with 0.5 mL·kg⁻¹·day⁻¹ of 0.01 M PBS as a negative control. The other two groups of ducks were also injected i.p. with either DHBcMAb or TAT PTD at 0.3 mg·kg^−1· day⁻¹ as DHBcMAb and TAT PTD controls respectively. The schedule of in vivo evaluations and assays is presented in Figure 2.

Figure 2.

In vivo assay schedule for the anti‐DHBV activity of DHBcMAb‐TAT PTD conjugate in ducks; d represents day.

Measurement of serum DHBV DNA by FQ‐PCR

The quantitative determination of serum DHBV DNA was performed using fluorescent quantitative (FQ)‐PCR, as described previously (Wang et al., 2013).

Determination of DHBV DNA and cccDNA in liver tissues by FQ‐PCR

Twenty to 50 mg of liver tissue was homogenized in 10 mM Tris–HCl (pH 7.5)‐10 mM EDTA‐0.5% SDS to a final concentration of 20 mg liver tissue mL⁻¹ using a ground‐glass homogenizer and then divided into two aliquots.

One aliquot, which was used to purify non‐protein‐bound cccDNA, was added to 5 M KCl to a final concentration of 0.5 M. After 30 min at room temperature, the mixture was centrifuged for 5 min at 12 000 g. The supernatant containing cccDNA was extracted twice with an equal volume of phenol saturated with 50 mM Tris–HCl (pH 8.0) and then extracted with an equal volume of phenol and chloroform (1:1) buffered with 500 mM Tris–HCl (pH 8). Nucleic acids were collected by ethanol precipitation at room temperature and resuspended in 10 mM Tris–HCl (pH 7.5)‐1 mM EDTA at 15 mg of liver tissue 40 μL⁻¹. To enhance the specificity of cccDNA detection, 500 ng of extracted nucleic acids was digested for 1 h in a final volume of 20 μL using 2 U Plasmid‐safe DNase (Epicentre, Madison, WI, USA) to degrade the relaxed‐circle and single‐stranded forms of the viral DNA, followed by DNase inactivation at 70°C for 30 min. Finally, 4 μL of the mixture was used for real‐time PCR amplification in an ABI Prism 7300 Sequence Detection System (Perkin‐Elmer Applied Biosystems, Foster City, CA, USA). The PCR mixture contained 12.5 μL 2 × HotStart Taq PCR MasterMix containing 0.1 U HotStart Taq Polymerase μL⁻¹, 500 μM of each dNTP, 20 mM Tris–HCl (pH 8.3), 100 mM KCl, 3 mM MgCl₂ (TIANGEN Biotech, Beijing, China), 1 μL (5 μM) of forward primer (5′‐GGCACAAACCTCCTGATT‐3′ nt 2445–2462), 1 μL (5 μM) of reverse primer (5′‐ACACATTGGCTAAGGCTC‐3′ nt 2666–2683), 1.25 μL (10 μM) of SyBr Green, 4 μL of template and 5.25 μL of diethylpyrocarbonate‐H₂O. After the initial denaturation at 94°C for 2 min, the thermal conditions consisted of 40 cycles of 94°C for 20 s, 49°C for 15 s, and 72°C for 30 s. Serial dilutions of a plasmid DHBV monomer (pBR322/2DHBV) were used as quantification standards. DHBV DNA was quantified using a standard curve. In this assay, the linear range was 1 × 10³–1 × 10⁸ copies mL⁻¹.

The other aliquot, which was used to purify total DNA, was digested with protease K for at least 2 h and then subjected to one phenol extraction and one phenol‐chloroform extraction. Precipitation was achieved with 2 volumes of ethanol in the presence of 0.3 M sodium acetate buffer (pH 5.2) followed by digestion with 100 μg of RNase mL⁻¹ in 10 mM Tris–HCl (pH 7.5)‐1 mM EDTA using 15 mg of liver tissue per 80 μL. Finally, 2 μL of the mixture was used for real‐time PCR amplification in an ABI Prism 7300 Sequence Detection System (Perkin‐Elmer Applied Biosystems, Foster City, CA, USA), as described for the quantification of DHBV DNA.

The inhibition ratios of each treatment for the levels of duck liver cccDNA were calculated using the following formula: inhibition ratio = (the level of cccDNA at baseline of treatment − the level of cccDNA at day 30 of treatment or at day 15 after the termination of treatment)/the level of cccDNA at treatment baseline.

Immunohistological examination of DHBcAg in the duck liver

Liver tissues fixed in formalin were dehydrated, embedded in paraffin, and sectioned to a thickness of 3 μm; then, the sections were deparaffinized and rehydrated. After the blocking endogenous peroxidases with 0.3% H₂O₂ for 10 min, non‐specific binding sites were blocked with 10% goat serum (ZSGB‐BIO Co., Beijing, China) for 10 min. Subsequently, the sections were incubated with a 1:50 dilution of mouse anti‐DHBV core MAb overnight at room temperature, followed by rabbit anti‐mouse biotin conjugate (ZSGB‐BIO Co., Beijing, China) for 30 min and HRP‐conjugated streptavidin (ZSGB‐BIO Co., Beijing, China) for 15 min. Immunostaining was developed using 3,3′‐diaminobenzidine (DAB) (ZSGB‐BIO Co., Beijing, China), and sections were counterstained with haematoxylin. Negative controls were performed by substituting the primary antibody with non‐specific immunoglobulins.

All immunoreactive areas of DHBcAg were quantified and analysed using Image‐Pro Plus 6.0 software (Media Cybernetics, Silver Spring, MD, USA) according to the manufacturer's instructions. For comparison of the mean density (MD), all fields were collected under the same photomicrography conditions simultaneously. The MD of each high‐power field (HPF) corresponding to the immunohistochemical positive degree of each slide was calculated as the integrated optical density of the positive areas divided by the measured areas of the fields. The MD of each immunohistochemical section was observed by counting the density of 10 randomly selected HPFs and averaging the results. The actual MD was calculated by subtracting the MD of the negative control from the tested section. The results are expressed as the mean ± SD. The MD of each duck at baseline was also subtracted from the MD at day 15 after treatment cessation, and the values were used to calculate the average value in each group. This average value was used for comparisons between groups.

Liver function, renal function and haematology determination

Duck blood samples were collected at each time point, and the serum levels of alanine aminotransferase (ALT), aspartate aminotransferase (AST), creatinine (CRE) and blood urea nitrogen (BUN) were analysed with an Olympus AU5400 automation chemistry system instrument (Olympus, Tokyo, Japan) using Olympus diagnostic kits. Hb was analysed with a Sysmex XT‐1800i automatic blood cell analyser (Sysmex, Kobe, Japan) using Sysmex diagnostic kits. Leukocytes (WBCs) and erythrocytes (RBCs) were analysed by manual counting with a microscope.

Data and statistical analyses

The data and statistical analyses comply with the recommendations for experimental design and analysis in pharmacology (Curtis et al., 2015). The randomized block design was applied for experimental grouping, and the experimental procedures or treatment and data analyses were conducted in a blinded fashion. All groups were designed to contain six ducks. For statistical analyses, we used raw data, except for those data on duck liver cccDNA quantification by FQ‐PCR, which were normalized to the inhibition ratios based on the cccDNA levels at the treatment baseline to reduce the effect of variable baselines. All statistical analyses were performed using SPSS software version 13.0 (SPSS, Chicago, IL, USA). Data were tested for normality using the Kolmogorov–Smirnov method. One‐way ANOVA followed by the least significant difference post hoc test were run if the F‐test of variance achieved P < 0.05, and there was no significant variance inhomogeneity, or the non‐parametric Mann–Whitney tests were run. Each sample was tested in triplicate for all experiments. Two‐sided P < 0.05 indicates a statistically significant difference.

Results

Binding of DHBcMAb‐TAT PTD conjugate to DHBcAg

The DHBcMAb‐TAT PTD conjugate was generated as described in the Methods section. To determine whether DHBcMAb and TAT were coupled successfully and confirm that the conjugate could bind DHBcAg, the binding of the DHBcMAb‐TAT PTD conjugate to DHBcAg was evaluated with ELISA using a DHBcAg precoated plate and mouse anti‐TAT‐HRP as the detection reagent. As the concentrations of the DHBcMAb‐TAT PTD conjugate and mouse anti‐TAT antibody decreased, the optical density gradually decreased in a dose‐dependent manner. In contrast, the optical densities of the DHBcMAb, TAT and negative control remained at the level of background values (Figure 1C).

Antibody titre detection

Antibody titres were determined based on P/N values (OD₄₅₀ test/OD₄₅₀ negative) exceeding 2.1. The DHBcMAb titres were 1:32 000, 1:64 000, and 1:16 000 after purification, concentration and coupling with TAT PTD respectively. These results suggest that DHBcMAb coupled with TAT PTD can recognize DHBcAg (Table 1).

Table 1.

DHBcMAb titres detected

Sample	OD450
	1:500	1:1000	1:2000	1:4000	1:8000	1:16 000	1:32 000	1:64 000	Negetive
DHBcMAb after purification (0.8 mg·mL−1)	2.15	1.92	1.12	0.58	0.25	0.11	0.07*	0.05	0.03
DHBcMAb after concentration (3.5 mg·mL−1)	3.86	2.40	1.79	1.03	0.65	0.31	0.21	0.13*	0.06
DHBcMAb‐TAT PTD (0.6 mg·mL−1)	3.12	2.22	1.45	0.61	0.38	0.17*	0.11	0.08	0.06

^* P/N(OD_{450 test}/OD_{450 negetive}) > 2.1

Inhibitory effect of the DHBcMAb‐TAT PTD conjugate on the levels of serum DHBV DNA in a duck model of HBV

DHBV‐infected ducks were treated i.p. with 0.03, 0.1 or 0.3 mg·kg⁻¹·day⁻¹ of the DHBcMAb‐TAT PTD conjugate or p.o. with 10.0 mg·kg⁻¹·day⁻¹ of adefovir dipivoxil for 30 days. Blood samples were collected at day 0, 10, 20 and 30 of treatment and at day 15 after the termination of treatment to quantify DHBV DNA. Liver samples were harvested at day 30 of treatment and at day 15 after the termination of treatment to quantify DHBV DNA and cccDNA (Figure 2).

At day 10 after the initiation of treatment, the serum DHBV DNA levels in the ducks treated with 0.3 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate were compared to those in the negative controls and those at the treatment baseline. The serum DHBV DNA levels at day 20 of treatment were significantly reduced compared to those of the negative controls and the treatment baseline, and those at day 30 of treatment were significantly reduced. Notably, the DHBV DNA levels remained significantly lower at day 15 after the termination of treatment than the levels of the negative controls and the treatment baseline (Figure 3). Similar inhibitory effects were observed for treatment with 0.1 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate. The inhibitory effect of 0.1 mg·kg⁻¹·day⁻¹ of DHBcAg antibody‐TAT conjugate was lower than that of 0.3 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT conjugate. The serum DHBV DNA levels in ducks treated with 0.03 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate decreased at day 10, 20 and 30 after the initiation of treatment; however, significant differences were only observed at day 30 compared to the levels in the negative controls and the treatment baseline. The inhibitory effects of 10.0 mg·kg⁻¹·day⁻¹ adefovir dipivoxil were similar to those of 0.3 mg·kg⁻¹·day⁻¹ DHBcMAb‐TAT conjugate, except that the levels of serum DHBV DNA at day 10 after the initiation of treatment were only significantly reduced by adefovir dipivoxil and not by 0.3 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT conjugate (Figure 3). This finding suggests that adefovir dipivoxil is a rapid‐acting DHBV inhibitor. Notably, at day 15 after the termination of treatment, the serum DHBV DNA levels were restored in all treatment groups; however, the levels of serum DHBV DNA in ducks treated with 0.3 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate remained significantly lower than those treated with adefovir dipivoxil.

Figure 3.

In vivo inhibitory effect of DHBcMAb‐TAT PTD conjugate on duck serum DHBV DNA levels. (A) Comparisons at the same time point. (B) Comparisons of the various treatments at different time points. NC, negative control; PC, positive control. Data are presented as the means ± SD (n = 6 per group). The common logarithm values of the concentrations of duck serum DHBV DNA were assessed using one‐way ANOVA. * P < 0.05.

DHBcMAb‐TAT PTD conjugate treatment dose‐dependently reduced the levels of DHBV DNA and cccDNA in the liver

To further evaluate the anti‐DHBV effect of the DHBcMAb‐TAT PTD conjugate in ducks, the levels of DHBV DNA and cccDNA in liver tissues were examined. At day 30 of treatment, the levels of liver DHBV DNA in ducks treated with 0.3 and 0.1 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate and 10 mg·kg⁻¹·day⁻¹ of adefovir dipivoxil were significantly reduced compared to the negative controls and the treatment baseline. However, the levels of liver DHBC DNA in those treated with 0.03 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate were comparable to the negative controls and the treatment baseline. At day 15 after the termination of treatment, only the levels of liver DHBV DNA in ducks treated with 0.3 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate were significantly reduced (Figure 4).

Figure 4.

In vivo inhibitory effect of DHBcMAb‐TAT PTD conjugate on duck liver DHBV DNA levels. (A) Comparisons at the same time point. (B) Comparisons of the PC and DHBcMAb‐TAT PTD (0.1 and 0.3 mg·kg⁻¹) treatments at different time points. NC, negative control; PC, positive control. Data are presented as the means ± SD (n = 6 per group). The common logarithm values of the levels of duck liver DHBV DNA were assessed using one‐way ANOVA. * P < 0.05.

At day 30 of treatment, the inhibition ratios of cccDNA in ducks treated with 0.3 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate and 10.0 mg·kg⁻¹·day⁻¹ of adefovir dipivoxil were significantly increased compared to the negative controls, whereas the inhibition ratios of cccDNA in ducks treated with 0.1 and 0.03 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate were comparable to the negative controls. At day 15 after the termination of treatment, the inhibition ratio of liver cccDNA was significantly increased only in the ducks treated with 0.3 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate compared to the negative controls (Figure 5).

Figure 5.

In vivo inhibitory effect of DHBcAg MAb‐TAT PTD conjugate on duck liver cccDNA levels. (A) Day 30 of treatment (end of treatment). (B) Day 15 after the termination of treatment. NC, negative control; PC, positive control. The inhibition ratios of each treatment on the level of duck liver cccDNA were calculated as described in the Methods section (n = 6 per group). The statistical analyses were carried out using a nonparametric Mann–Whitney test. * P < 0.05.

These data show that, consistent with its inhibitory effect on the serum DHBV DNA level, DHBcMAb‐TAT PTD conjugate treatment reduced the levels of DHBV DNA and cccDNA in the liver in a dose‐dependent manner. Unlike adefovir dipivoxil, the anti‐DHBV effect of 0.3 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate persisted for at least 15 days after the termination of treatment.

Immunohistological analyses of DHBcAg were performed in livers from various treatment groups at the treatment baseline and at day 15 after the termination of treatment. At baseline, DHBcAg was detected in the cytoplasm of hepatocytes (Figure 6A). At day 15 after the termination of treatment, the DHBcAg expression levels in ducks treated with 0.3 and 0.1 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate were lower than those in the negative controls and at baseline. In contrast, the DHBcAg expression levels in ducks treated with 0.03 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate and 10 mg·kg⁻¹·day⁻¹ of adefovir dipivoxil were comparable to those of negative controls and the treatment baseline (Figure 6A,B). Differences between the MDs at the treatment baseline and the end of treatment in the ducks that received 0.3 and 0.1 mg·kg⁻¹·day⁻¹ of DHBcMAb‐TAT PTD conjugate were significantly higher than those in the negative controls (Figure 6B).

Figure 6.

Immunohistological analysis of DHBcAg in duck liver sections. (A) Liver tissue in each DHBV infected duck in all groups was obtained at day 10 before initiation of treatment (baseline of treatment) and at day 15 after the termination of treatment. Liver sections were subjected to immunohistological analysis of DHBcAg. Representative photographs from the same duck in each group are presented (magnification: 600×). (B) Mean density of the DHBcAg staining in duck liver tissues was calculated using the Image‐Pro Plus 6.0 software. The differences between mean density at baseline and at end of treatment are presented as the means ± SD (n = 6 per group) and comparisons between groups were assessed by one‐way ANOVA. NC, negative control; PC, positive control. * P < 0.05.

Notably, DHBcMAb‐TAT PTD conjugate treatment exhibited no significant toxicity in the animals, because all ducks in the treated and control groups grew equally well, and no significant weight change or abnormal behaviour was observed during treatment. Furthermore, no significant changes between baseline and end of treatment were observed in ALT, AST, CRE, BUN, WBCs, RBCs and Hb in the ducks from the various treatment groups, including the different DHBcMAb‐TAT PTD conjugate dosing groups (Supporting Information).

Discussion

Over the past decade, interest in the use of antibodies against intracellular targets has been growing. An intrabody, which is an antibody or fragment of an antibody that is expressed intracellularly and can be directed to a specific target antigen present in various subcellular locations, is a novel tool for the management of viral infections (Marasco, 1995; Wheeler et al., 2003; Boons et al., 2014; Suzuki et al., 2016). In particular, intrabodies targeting HBcAg have been reported in two studies (Yamamoto et al., 1999; Serruys et al., 2010). In one of these studies, HBcAg‐specific single‐chain antibodies (scFvs) efficiently shut down viral DNA replication (Yamamoto et al., 1999). In the other study, HBcAg‐specific single‐domain antibodies (VHH) targeted to the nucleus affected HBcAg and HBeAg expression (Serruys et al., 2010).

A transbody, which is a ‘cell‐permeable’ antibody constructed by coupling PTD to antibodies, differs from the conventional intrabodies expressed within cells. Indeed, transbodies represent a new strategy for using antibodies against intracellular targets. Designs of transbodies using scFvs (Poungpair et al., 2010) or humanized single‐domain antibodies (Thueng‐in et al., 2012; Phalaphol et al., 2013; Glab‐Ampai et al., 2016) have been reported. In our previous study, the HBV transbody, which was prepared by coupling HBcMAb to TAT PTD, was determined to possess cell‐penetrating ability and potent antiviral activity in vitro.

In this study, DHBV replication in ducks was used to evaluate the antiviral activity of DHBcMAb‐TAT PTD conjugate in vivo. DHBV is a member of the hepadnaviridae family that shares similarities with human HBV in terms of its genome structure, virus replication strategy and outcomes of infection (Jilbert et al., 1996; Jilbert et al., 1998; Bertoletti and Ferrari, 2003; Foster et al., 2005). DHBV‐infected ducks represent a useful model for testing novel antiviral approaches for human HBV infection (Wu et al., 2007; Noordeen et al., 2013; Tang et al., 2013). In addition, unlike previous studies examining antiviral activity using transgenic animals (Serruys et al., 2009; Poungpair et al., 2010; Thueng‐in et al., 2012), the present study used a naturally infected host, DHBV‐infected ducks, to investigate the antiviral activity of the transbody. This model is more clinically relevant because it reflects the intrinsic features of viral infection and the actual activities of the transbody.

Examining the antiviral activity of the DHBV transbody in DHBV‐infected ducks demonstrated that this transbody has potent anti‐DHBV activity in vivo. Administering the DHBV transbody for 30 days significantly reduced the serum and liver DHBV DNA levels at doses of 0.1 and 0.3 mg·kg⁻¹·day⁻¹. In addition, when the DHBV transbody was administered at 0.3 mg·kg⁻¹·day⁻¹ for 30 days, the cccDNA levels in the liver decreased significantly. Notably, the suppressive effects of 0.3 mg·kg⁻¹·day⁻¹ DHBV transbody on the serum and liver DHBV DNA and liver cccDNA levels remained significant 15 days after treatment cessation, indicating the relatively long‐lasting in vivo anti‐HBV effect of this transbody. Immunohistochemistry of liver sections also revealed decreased DHBcAg within the hepatocytes at day 15 after treatment termination in ducks administered 0.1 and 0.3 mg·kg⁻¹·day⁻¹ of the transbody. This finding further supports the long‐lasting activity of the DHBcMAb‐TAT PTD conjugate in suppressing virus replication.

These findings suggest that the DHBcMAb‐TAT PTD conjugate, a cell‐permeable antibody or transbody, retained the correct conformational folding and disulfide bond formation in the reducing conditions within cells, which is a distinct advantage over conventional intrabodies expressed within cells. For intrabodies, the initial conformational folding and disulfide bond formation are adversely affected by the reducing conditions within cells (Wörn and Plückthun, 2001). More importantly, the use of a cell‐permeable antibody would avoid the safety and ethical concerns associated with the direct application of recombinant DNA technology in human clinical therapy, because the intrabody must be expressed within cells (Heng and Cao, 2005).

Although the exact mechanism by which the DHBV transbody inhibits DHBV replication requires further study, the interaction between the DHBV transbody and HBcAg in cells is undoubtedly a decisive factor. Combined with the results of our previous study (Wang et al., 2015), the transbody against HBcAg can interfere with two major steps in the HBV life cycle: nucleocapsid assembly and HBV DNA transcription. HBcAg encapsidates the pregenome RNA (pgRNA), reverse transcriptase and host factors to form the nucleocapsid, which provides the structural background for pgRNA reverse transcription into viral DNA (Bruss, 2004; 2007). Moreover, HBcAg is an important component of the cccDNA minichromosome and preferentially binds to the CpG island 2 of HBV cccDNA and serves as a positive regulator of HBV transcription and replication (Bock et al., 2001; Vivekanandan et al., 2009; Guo et al., 2011).

The HBV replication cycle involves multiple complex steps, which occur in both the cytosol and nucleus of infected cells. To study the mode of action of the transbody, HBV DNA‐integrated hepatoma cells (HepG2.2.15 cells) were treated with the HBcMAb‐TAT PTD conjugate. The intracellular localization of the HBcMAb‐TAT PTD conjugate was determined using mouse anti‐TAT‐fluorescein isothiocyanate (FITC). As shown in Supporting Information Fig. S8, the fluorescence signal was detected in both the cytoplasm and nucleus of the cells, indicating the localization of the transbody in both the cytoplasm and nucleus. Additionally, the levels of HBcAg were reduced in a dose‐dependent manner. HBcAg plays an important role in nucleocapsid assembly. Because nucleocapsid assembly mainly occurs in the cytoplasm, the localization of transbodies against HBcAg in the cytosol of infected cells and the reduction of HBcAg suggest that it may interfere with nucleocapsid assembly. To study the direct effect of the transbody on the transcription of HBV DNA, HepG2.2.15 cells, which produce infectious viral particles, were treated with various concentrations of the HBcMAb‐TAT PTD conjugate, and the levels of extracellular (secreted) and intracellular HBV DNA were measured. As shown in Supporting Information Fig. S9, the HBcMAb‐TAT PTD conjugate dose‐dependently inhibited HBV DNA production and secretion. Taken together, these results show that the transbodies prepared in this study may interfere with both nucleocapsid assembly and HBV DNA transcription. However, the main purpose of this study was to evaluate the antiviral activity of the DHBcMAb‐TAT PTD conjugate in vivo, and determining the mechanism by which the transbody affects the HBV life cycle in more detail requires further study.

In addition, no significant changes in the ALT and AST were observed between before and after treatment in ducks administered the DHBcMAb‐TAT PTD conjugate, suggesting that a much longer duration of DHBcMAb‐TAT PTD conjugate treatment may be required to observe reduced ALT and AST. In vivo administration of the DHBcMAb‐TAT PTD conjugate exhibited no significant toxicity in the ducks. This finding is important for the long‐term treatment of HBV infection.

Overall, the present study demonstrated that the DHBcMAb‐TAT PTD conjugate has potent antiviral activities in vivo. This cell‐permeable antibody or transbody against HBcAg may provide a novel approach for the treatment of HBV infection in humans. The effects of the HBcMAb‐TAT PTD conjugate on nucleoside analogue‐resistant HBV and different HBV genotypes and of co‐treatment with the HBcMAb‐TAT PTD conjugate and a nucleoside analogue warrant further investigation.

Author contributions

Y.L., Z.L. and Y.W. conceived and designed the experiments. Y.L., L.H., X.L., A.F., W.W., L.Z., N.L. and Y.W. performed the experiments. G.Z., Q.W. and Q.H. analysed the data. Y.L. and Y.W. drafted the manuscript. Y.L., Q.W. and G.Y. revised the paper. Z.L. and Y.W. supervised the project.

Conflict of interest

The authors declare no conflicts of interest.

Declaration of transparency and scientific rigour

This Declaration acknowledges that this paper adheres to the principles for transparent reporting and scientific rigour of preclinical research recommended by funding agencies, publishers and other organisations engaged with supporting research.

Supporting information

Figure S1 Serum alanine aminotransferase (ALT) in each group at each time point (n = 6 per group). There is no significant change among each group at each time point (P > 0.05).

Figure S2 Serum aspartate aminotransferase (AST) in each group at each time point (n = 6 per group). There is no significant change among each group at each time point (P > 0.05).

Figure S3 Serum creatinine (CRE) in each group at each time point (n = 6 per group). There is no significant change among each group at each time point (P > 0.05).

Figure S4 Serum blood urea nitrogen (BUN) in each group at each time point (n = 6 per group). There is no significant change among each group at each time point (P > 0.05).

Figure S5 Hemoglobin (HGB) in each group at each time point (n = 6 per group). There is no significant change among each group at each time point (P > 0.05).

Figure S6 Leukocyte (WBC) in each group at each time point (n = 6 per group). There is no significant change among each group at each time point (P > 0.05).

Figure S7 Erythrocytes (RBC) in each group at each time point (n = 6 per group). There is no significant change among each group at each time point (P > 0.05).

Figure S8 Intracellular localization of the HBcMAb–TAT PTD conjugate in HepG2.2.15 cells and the inhibitory effect of HBcMAb–TAT PTD conjugate on HBcAg levels. (A) HepG2.2.15 cells treated with 40.0 μg mL⁻¹ of HBcMAb–TAT PTD conjugate. (B) An enlargement of the white‐frame area in (A). (C) HepG2.2.15 cells treated with 40.0 μg mL⁻¹ of HBcMAb. (D) inhibition ratio = (the level of HBcAg treated with PBS – the level of HBcAg treated with HBcMAb, BSA‐TAT PTD or the HBcMAb–TAT PTD conjugate) / the level of HBcAg treated with PBS. (n = 3). * P < 0.05, significantly different from the inhibition ratio treated with HBcMAb. # P < 0.05, significantly different from the inhibition ratio treated with BSA–TAT PTD.

Figure S9 The inhibitory effect of HBcMAb–TAT PTD conjugate on HBV DNA levels. (A) in cell lysates. (B) in cell culture medium. (n = 3). inhibition ratio = (the level of HBV DNA treated with PBS – the level of HBV DNA treated with HBcMAb, BSA‐TAT PTD or the HBcMAb–TAT PTD conjugate) / the level of HBV DNA treated with PBS. * P < 0.05, significantly different from the inhibition ratio treated with HBcMAb. # P < 0.05, significantly different from the inhibition ratio treated with BSA–TAT PTD.

Li, Y. , Liu, Z. , Hui, L. , Liu, X. , Feng, A. , Wang, W. , Zhang, L. , Li, N. , Zhou, G. , Wang, Q. , Han, Q. , Lv, Y. , Wang, Q. , Yang, G. , and Wang, Y. (2017) Transbody against virus core protein potently inhibits hepadnavirus replication in vivo: evidence from a duck model of hepatitis B virus. British Journal of Pharmacology, 174: 2261–2272. doi: 10.1111/bph.13811.