Chimeric monoclonal antibodies to hypervariable region 1 of hepatitis C virus

Chengyao Li1 and Jean-Pierre Allain2


1 National Blood Service, Division of Transfusion Medicine, East Anglia Blood Centre, Long Road, Cambridge CB2 2PT, UK
2 Department of Haematology, Division of Transfusion Medicine, East Anglia Blood Centre, Long Road, Cambridge CB2 2PT, UK

Correspondence
Jean-Pierre Allain
jpa1000{at}cam.ac.uk


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Two chimeric monoclonal antibodies (cAbs), 2P24 and 15H4, to hypervariable region 1 (HVR1) of hepatitis C virus (HCV) were constructed by grafting the variable regions of murine monoclonal antibodies (mAbs) 2P24 and 15H4 to a human IgG1 kappa constant region. Two cAb-producing cell lines were adapted to serum-free media. Both cAb 2P24 and cAb 15H4 cell lines produced 3–5 µg antibodies ml–1 after 3–5 days culture. cAbs retained binding characteristics similar to those observed in the original mAbs. There was no clear difference in affinity between binding of cAbs and mAbs to seven HVR1 peptides. Mixtures of biotinylated cAbs or mAbs reacted with 32 (86 %) and 31 (84 %) of 37 HVR1 peptides, respectively, but not with non-HVR1 control peptides. HCV from 16 out of 18 (89 %) random HCV-containing plasmas was captured by the mixture of biotinylated cAbs. The capture from IgG-depleted plasmas suggested that cAbs captured mainly free rather than complexed HCV, irrespective of genotype. A mixture of the two cAbs inhibited HCV binding to Molt-4 cells in a dose-dependent manner. These cAbs may be useful for prevention of nosocomial HCV infection and passive immunization to prevent HCV reinfection after liver transplantation.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
An estimated 170 million people are infected with hepatitis C virus (HCV) worldwide (World Health Organization, 1999) and at least 3–4 million people are infected each year (Crabb, 2001). Over 70 % of acute infections eventually become persistent, of which a significant proportion develop liver cirrhosis and ultimately hepatocellular carcinoma (Crabb, 2001; Lavanchy & McMahon, 2000). At present, antiviral therapy with a combination of interferon and ribavirin is commonly used clinically with limited efficacy (Fried & Hoofnagle, 1995; McHutchison et al., 1998). The development of effective therapeutic drugs is essential for treatment and prevention of chronic hepatitis C.

Hypervariable region 1 (HVR1), a sequence of 27 residues at the N terminus of the main envelope protein E2 of HCV, is a target for neutralizing antibodies (Farci et al., 1994, 1996; Rosa et al., 1996; van Doorn et al., 1995) and a possible ligand of HCV binding to cells (Basu et al., 2004; Hamaia et al., 2001; Kurihara et al., 2004; Penin et al., 2001; Scarselli et al., 2002). However, HVR1 is highly mutated, which allows HCV to escape the host's immunity (Farci et al., 1996; Korenaga et al., 2001; Kumar et al., 1994; Ray et al., 1999; Shimizu et al., 1994). In recent years, a number of investigators have observed that antibodies from HCV-infected patients, or from mice and rabbits immunized with HVR1 peptides, cross-react with a wide range of HVR1 peptides and can be used to capture HCV variants and inhibit HCV binding to cells (Esumi et al., 1998; Mondelli et al., 1999; Puntoriero et al., 1998; Shang et al., 1999; Watanabe et al., 1999; Zibert et al., 1995). Monoclonal antibodies (mAbs) are a potent treatment against many infectious agents (Casadevall et al., 2004). As HVR1 is highly heterogeneous in its primary sequence, to obtain a mAb broadly recognizing HCV would appear difficult. Currently, most mAbs to HVR1 are poorly cross-reactive with HVR1 variants and have limited ability to recognize multiple HCV strains (Allander et al., 2000; Cerino et al., 2001; Triyatni et al., 2002; Zhou et al., 2000).

The specific immunotherapy and prevention of HCV infection by using broadly cross-reactive antibodies to HVR1 of HCV might be effective in clinical applications. In a previous study (Li et al., 2001), we discovered that a conserved epitope based on the G--Q motif at positions 23–26 within HVR1 was critical to induce cross-reactive antibodies to HVR1 variants. Two high-affinity murine mAbs to the G--Q epitope of HVR1, cross-reacting with 87 % of HVR1 peptides and highly effective in capturing HCV strains and blocking HCV binding to cells, were obtained. In order to avoid the antigenicity of murine mAbs in humans, chimeric monoclonal antibodies (cAbs) keeping only the murine variable regions were produced and characterized in comparison with the parental murine mAbs to HVR1 of HCV.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
HCV peptides.
A series of 37 synthetic peptides, each of 16–19 residues, corresponding to C-terminal sequences of HVR1 (covering 27 aa from positions 384 to 410 of the HCV polyprotein) and other HCV regions (core, E1, E2 and NS3) was obtained from Severn Biotech or Cambridge Research Biochemicals (see Table 2). Four 9mer HVR1 peptides (covering positions 19–27) were obtained from Severn Biotech: MH2-C (LFDLGPKQK), MH5-C (MFSLGARQK), G1245-C (LFNLGPQQQ) and EH-C (LFTPGAKQN). The purity of the peptides was above 75 %.


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Table 2. Cross-reactivity of chimeric antibodies to HVR1 peptides

 
HCV-positive samples.
Eight HCV-containing plasmas or sera (designated with the prefix UK) were collected from British blood donors and patients infected with HCV genotype 1, 2 or 3 at the East Anglia Blood Centre or from patients at Addenbrooke's Hospital, Cambridge, UK. Four samples (prefix G) were collected from Ghanaian blood donors infected with HCV genotype 2 at the Komfo Anokye Teaching Hospital blood bank, Kumasi, Ghana (Candotti et al., 2003). One sample (prefix US) was collected from an American patient infected with HCV genotype 1 and five samples (prefix X) were collected from chimpanzees infected chronically with HCV genotype 1 from American patients (kindly provided by Dr H. J. Alter, NIH, Bethesda, MD, USA). All samples contained antibodies to HCV and HCV RNA, detected by RT-PCR as described previously (Petrik et al., 1997).

Cloning of mAb variable-region genes.
The variable-region genes of the heavy (VH) and light kappa (VK) chains of mAbs 2P24 and 15H4 (Li et al., 2001) were amplified by using the mouse primers VH1 (forward, 5'-GGAACCCTTTGGCCCAGCCGGCCATGGCCSAGGTYCAGCTBCAGCAGTC-3') and CH (reverse, 5'-TARCCYTTGACMAGGCATCC-3'), and VK1 (forward, 5'-TATTCGTCGACGGATATTGTGATGACBCAGDC-3') and CK (reverse, 5'-CGTTCACTGCCATCAATC-3'), obtained from Dr T. Grunwald (Medical Research Council, Cambridge, UK), and the primers CK and MKS11 (forward, 5'-GCCCAGTTCCTGTTTCTG-3') (for the 2P24 VK chain only) obtained from Dr I. Harmer (Division of Transfusion Medicine, University of Cambridge, UK). The PCR products were cloned and sequenced as described previously (Li et al., 2001). The nucleotide and deduced amino acid sequences were analysed and defined by the Kabat numbering system (Kabat et al., 1991).

Construction of genes for cAbs.
The Kabat-numbered variable-region sequences of the VH and VK chains of mAbs 2P24 and 15H4 were isolated and modified by overlapping-extension PCR with Pwo DNA polymerase (Roche). The VH and VK DNA fragments were cloned into pSVgpt-B2VH-hucIgG1 and pSVhygFog-1V{kappa}-HuCK vectors with HindIII and BamHI sites by replacement of the V regions, respectively. These two vectors (Furtado et al., 2002) were kindly provided by Dr K. Armour (Department of Pathology, University of Cambridge, UK).

Transfection and antibody production.
DNA constructs containing the VH or VK chain of cAbs 2P24 and 15H4 were prepared by using an EndoFree Plasmid Maxi kit (Qiagen). The PvuII-linearized VH chain construct DNA (10 µg) was mixed with 20 µg PvuII-linearized VK chain construct DNA and co-transfected into 1x107 SP2/0 myeloma cells by electroporation (0·4 cm gap Bio-Rad cuvette, 180 V and 960 µF). After electroporation, the cells were added immediately to 20 ml RPMI 1640 medium containing 10 % FCS in a 75 cm2 flask and incubated for 48–72 h at 37 °C. The cells were resuspended in 40 ml RPMI 1640 medium with 10 % FCS containing 0·8 µg mycophenolic acid ml–1 and 250 µg xanthine ml–1 and distributed over two or three 96-well plates at 200 µl per well. After 3 weeks, the wells containing cell colonies were screened for the presence of cAb in the culture supernatant by ELISA.

Transfectomas producing cAb 2P24 or 15H4 were cloned repeatedly in RPMI 1640 medium containing 15 % FCS and 5 % BM Condimed H1 supplement (Roche). Clones were selected according to antibody-expression levels and stability. Several clones for each cAb were adapted to hybridoma serum-free medium (SFM; Gibco), serum-free and protein-free medium (SPF; Sigma) or a mixture of both SFM and SPF. Cells were passaged in the media for 3–5 days and supernatants were collected for antibody-level tests.

cAbs were purified from the supernatant of SFM or SFP cultures in flasks by using Protein G columns. Some antibodies were produced by MiniPerm (Vivascience) and purified by Protein G columns.

Antibody biotinylation.
cAbs and mAbs were biotinylated by using a Micro-Biotinylation kit (Sigma). Biotinylated antibodies were used to assess the ability to cross-react with HVR1 peptides or to capture HCV.

Enzyme immunoassay (EIA).
Indirect ELISA was used for the measurement of cAb-expression levels in the supernatants of cell cultures. Goat anti-human IgG Fc fragment-specific antibody (Sigma) was coated on the plates. Goat anti-human IgG{kappa}–alkaline phosphatase conjugate was used as a secondary antibody for detecting cAbs bound to the coated anti-human IgG Fc fragment. Human IgG1{kappa} (Sigma) was used as a standard.

The reactivity of biotinylated cAbs and mAbs with various HCV peptides was measured by peptide EIA in Nunc-Immuno plates (Maxisorp; Nalge Nunc) as described previously (Li et al., 2001). Levels of antibody reactivity to HVR1 peptides by EIA were presented as sample/cut-off (S/CO) ratios. The cut-off was calculated as the mean of the A405 values of non-HVR1 peptides+6SD.

Affinity measurements.
The affinity of the cAbs and mAbs was determined against seven selected keyhole limpet haemocyanin (KLH)/BSA-conjugated HVR1 peptides (EH, MH2, MH5, S67, S85 S90 and L1.1) (Jackson et al., 1997) by using an IAsys optical biosensor (Affinity Sensor) as described previously (Li et al., 2001; Zhai et al., 1999). Affinity constants (Kd) were calculated from these measurements as Kdiss/Kass by using the FASTFIT program.

Real-time quantitative RT-PCR analysis of HCV RNA.
HCV RNA was measured by real-time quantitative RT-PCR using the Mx4000 Multiplex Quantitative PCR system (Stratagene) as described previously (Candotti et al., 2003). For each run, duplicates of a tenfold serial dilution of WHO International Standard for HCV RNA for NAT assays 96 and 790 (NIBSC) containing 4x102–4x105 IU HCV genome ml–1 were used as a standard curve for quantification of HCV RNA. Each sample was analysed in duplicate and the results were averaged. The sensitivity of quantitative RT-PCR for detecting HCV in plasma was 100 IU ml–1.

HCV capture.
Unselected plasmas from chronically HCV-infected patients were centrifuged for 2 min at room temperature. The plasma supernatant was collected for two sample preparations: (i) plasma or its dilution (1 : 10 in PBS) (native sample) was untreated and used directly for the HCV capture test; (ii) plasma or its dilution (1 : 5 in PBS) was chromatographed through a 1 ml Protein G column (Amersham Biosciences) to obtain IgG-depleted HCV plasma for use in the HCV capture test. Native or IgG-depleted HCV plasma (100 µl) was added to a 50 µl mixture of biotinylated cAbs 2P24 and 15H4 or mAbs 2P24 and 15H4 in PBS (20 µg ml–1) containing 0·1 % Tween 20 and 4 % BSA and pre-incubated at 37 °C for 1·5 h and then overnight at 4 °C. Fifty microlitres of 2 mg streptavidin-coated magnetic particles ml–1 (Promega) was added to the biotinylated antibody–HCV mixture and incubated for 1 h at room temperature. Biotinylated mouse myeloma IgG1 (Sigma) was used as a negative control in each assay. After four washes with PBS containing 0·1 % Tween 20, HCV RNA was extracted from the magnetic particles by using a High Pure Viral RNA kit (Roche) and detected by real-time quantitative RT-PCR (Candotti et al., 2003).

Inhibition of HCV binding to target cells.
One hundred microlitres of various dilutions of cAbs and normal mouse myeloma IgG1 was pre-incubated with 50 µl IgG-depleted HCV-containing plasma or native HCV-containing plasma for 2 h at 37 °C and then at 4 °C overnight. The mixture was added to 2x105 Molt-4 cells in a 250 µl final volume and incubated at room temperature for 1 h. The cells were washed four times, and viral and cellular RNA was extracted by using an RNeasy Mini kit (Qiagen) and tested for the presence of HCV RNA by real-time quantitative RT-PCR as described above. Under identical conditions, 50 µl HCV-containing plasma without cAb pre-incubation and normal plasma were added to cells as positive and negative controls, respectively.


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Testing of the variable-region sequences of mAbs 2P24 and 15H4
The variable-region genes of mAbs 2P24 and 15H4 were isolated and modified for preparing the cAb constructs. The amino acid sequences of the variable regions of the VH and VK chains were numbered by the Kabat numbering system. The VH and VK chains of mAb 15H4 contained 118 and 114 aa, respectively, and those of mAb 2P24 contained 114 and 115 aa, respectively. At position 69 of the 2P24 VH nucleotide sequence, the original A was mutated to G to eliminate the HindIII restriction site for cloning the VH fragment into the expression vector using HindIII/BamHI sites, but the cDNA-encoded amino acid sequence was unchanged.

Production of cAbs
Two human IgG1 versions of mAbs 2P24 and 15H4 were produced. Chimeras of VK and VH were made, retaining the original murine complementarity-determining regions and framework regions, but with the human CK and CH1 (IgG1) constant regions, respectively. In the cAb-expressing vectors (Furtado et al., 2002), the complete DNA sequence of cAb 2P24 or 15H4 contained, at its 5' end, the Ig promoter, the eukaryotic exon and intron leader sequences, the last 4 aa of the leader region (GVHS, which form part of the V region exon) and, at its 3' end, the 5' end of the first half of the V–C intron.

After co-transfection of SP2/0 with the VH and VK chain construct DNAs containing murine mAb variable regions and human antibody (IgG1{kappa}) constant regions, the full-length cAb VH and VK chains were expressed and whole molecules of cAb were assembled and secreted in the culture medium. The cAb 2P24-producing cell line was cloned seven times and adapted to a mixture of 25 % SFM and 75 % SPF. The cAb 15H4 cell line was cloned three times and adapted to SFM. The final clones of the cAb 2P24 cell line in SFM/SPF medium and the cAb 15H4 cell line in SFM were passaged for more than 6 months and remained stable. Levels of cAbs 2P24 and 15H4 in the supernatants ranged from 2 to 4 µg ml–1 and from 3 to 5 µg ml–1, respectively, after 3–5 days in culture. The concentrations of cAbs 2P24 and 15H4 reached 10 µg ml–1 in the saturated cell-culture flasks and 36 µg ml–1 in MiniPerm (Vivascience) culture.

Fine mapping of the conserved HVR1 epitope recognized by the cAbs
Our previous data suggested that mAbs 2P24 and 15H4 recognize a G--Q-based motif of HCV HVR1, as HVR1 peptide substituted at G and Q (positions 23 and 26 of the HVR1 sequence) with V and L, respectively, no longer reacts with the mAbs (Li et al., 2001). To confirm that the basic structure of G--Q was a conserved epitope, four 9mer HVR1 peptides, MH2-C, MH5-C, G1245-C and EH-C (covering positions 19–27), and a 15mer, MH2, containing the G--Q motif with substitutions at positions other than positions 23 and 26, were tested in a competitive EIA. Fig. 1(a) shows that the binding of cAb 2P24 to the 15mer HVR1 peptide MH2 was inhibited competitively by 9mer HVR1 peptides MH2-C, MH5-C, G1245-C, EH-C and MH2, but not by a control core peptide, S5. In contrast, the binding of cAb 15H4 to MH2 was not affected by any of the 9mer HVR1 peptides or by S5, but was inhibited competitively by the 15mer HVR1 peptide, MH2 (Fig. 1b). The results further confirmed that the recognition of cAb 2P24 was based entirely on the conserved G--Q motif and that the recognition of cAb 15H4 was not limited to that motif. The recognition of the conserved HVR1 epitopes by cAbs 2P24 and 15H4 was thus maintained from the original murine mAbs 2P24 and 15H4.



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Fig. 1. Competitive inhibition of cAb binding to HVR1 by 9mer HVR1 peptides. Biotinylated cAb 2P24 (a) or 15H4 (b) (0·5 µg ml–1) was mixed with the four 9mer HVR1 peptides MH2-C ({blacklozenge}), MH5-C ({blacksquare}), EH-C ({blacktriangleup}) and G1245-C (x) at concentrations of 0, 1, 10 and 100 µg ml–1 and incubated for 1 h at 37 °C. MH2 HVR1 (*) and S5 ({bullet}) core peptides were used as positive- and negative-inhibition controls, respectively. Biotinylated mouse myeloma IgG1 (N-mAb, +) was used as an antibody-negative control. The mixture (100 µl) was added to MH2 HVR1 peptide-coated wells and incubated for 1 h at 37 °C. Bound biotinylated cAb was detected by streptavidin–alkaline phosphatase by using para-nitrophenyl phosphate substrate.

 
Comparison of the affinity of cAbs and mAbs for HVR1 peptides
Seven KLH/BSA-coupled HVR1 peptides were immobilized on the activated surface of carboxymethyl dextran cuvettes and used to measure the affinity of cAbs and mAbs using an IAsys optical biosensor. HVR1 peptides EH, MH2, MH5, S67, S85 and S90 reacted with both cAbs (S85 did not react with cAb 15H4) and mAbs. Peptide L1.1 reacted weakly with cAbs and mAbs. The affinity constants (Kd) and EIA values are presented in Table 1. The affinity of cAbs and mAbs was similar except for cAb 15H4, which did not react with S85. The affinity of cAbs for some HVR1 peptides was slightly higher than the equivalent mAb. The affinity of cAb 2P24 to six EIA-reacting HVR1 peptides and cAb 15H4 to five EIA-reacting peptides reached 10–8 to 10–9 M.


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Table 1. Comparison of the binding affinity of cAbs and mAbs to HVR1 peptides

 
Comparison of cross-reactivity of cAbs and mAbs with HVR1 peptides
Thirty-seven HVR1 peptides and five non-HVR1 HCV peptides were used to test the cross-reactivity and specificity of the cAbs. The reactivity of a mixture of biotinylated cAbs 2P24 and 15H4 and of a mixture of biotinylated mAbs 2P24 and 15H4 to peptides was measured by EIA and the results are presented in Table 2. cAbs and mAbs reacted with 32 (86 %) and 31 (84 %) out of 37 HVR1 peptides, respectively, but not with control peptides. The reactivity of cAbs and mAbs was similar, but S/CO levels showed slight discrepancies for some HVR1 peptides in both directions. A decrease in reactivity of cAb 15H4 with some peptides was observed, but this relative defect was compensated for by cAb 2P24.

HCV capture by cAbs and mAbs
Eighteen non-selected HCV-containing plasmas were used for HCV capture by the cAb or mAb mixture. HCV was captured from native plasmas that contained antibody–HCV complexes and from IgG-depleted plasmas that contained mostly uncomplexed virus. The captured HCV was detected by real-time RT-PCR. Eighty-nine per cent of HCV strains were captured (Table 3). Results suggested that the antibody capacity for HCV capture was not dependent on the genotype of HCV and that mostly free HCV was captured. The first round of HCV capture from IgG-depleted plasmas ranged from 1 to 60 % of total HCV, but complexed virus in native plasma was captured poorly. The ability to capture more HCV in a second round of capture of the unretained (free) virus fraction suggested that the majority of free virus was susceptible to capture (data not shown).


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Table 3. Capture of HCV by biotinylated antibodies to HVR1

Viral loads and captured HCV were measured by quantitative RT-PCR and are indicated in IU ml–1. ND, Not determined.

 
Ability of cAbs to inhibit HCV binding to target cells
A mixture of cAbs 2P24 and 15H4 or a control mouse myeloma IgG1 was pre-incubated with UKS3 (genotype 3a) native and UKS2 (genotype 1a) IgG-depleted HCV plasmas and added to Molt-4 cells. Bound HCV was detected by real-time RT-PCR. The results showed that 0·8 µg cAbs ml–1 inhibited 55·2 and 78·3 % of binding of UKS2 IgG-depleted or UKS3 native HCV to Molt-4 cells, respectively (Fig. 2a), and that this inhibition was antibody dose-dependent (Fig. 2b).



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Fig. 2. Inhibition of HCV binding to Molt-4 cells by cAbs. (a) IgG-depleted UKS2 (2x105 IU ml–1) and native UKS3 (7·4x105 IU ml–1) HCV plasmas were pre-incubated separately with 0·8 µg cAb 2P24 and 15H4 mixture ml–1 (filled bars) or a mouse myeloma IgG1 control (shaded bars) and then added to 2x105 Molt-4 cells. (b) Native UKS3 HCV plasma (7·4x105 IU ml–1) was pre-incubated with a mixture of cAbs 2P24 and 15H4 at increasing concentrations and then added to Molt-4 cells. Bound HCV was detected by quantitative RT-PCR and is given in IU ml–1. Data are presented as percentage inhibition and are means±SD of values from three independent experiments.

 

   DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
The therapeutic use of antibodies can be traced back more than a century, when mice were first investigated as a potential source. By injecting mice with infectious agents, scientists aimed to stimulate the production of antibodies targeted against the infection. Köhler & Milstein (1975) invented hybridoma technology, which allowed researchers to mass-produce individual antibodies for the first time. In order to reduce murine antigenicity of mAbs, chimeric (Bruggemann et al., 1987) and fully humanized (Jones et al., 1986) antibodies were created by genetic engineering. To date, the US Food and Drug Administration has approved more than 12 therapeutic mAbs, most of them in the past 5 years (Casadevall et al., 2004). Five are cAbs (Gura, 2002). Most cAbs are suitable for clinical use by maintaining the original antibody capacity, with less or no immunogenicity in humans. By comparison, fully humanized antibodies reduce immunogenicity in humans maximally, but frequently lose affinity (Clark, 2000). In this study, we generated two cAbs containing the variable regions of two murine IgG1{kappa} mAbs and the constant regions of a human IgG1{kappa} antibody.

Two cloned transfectomas harbouring cAb 2P24- or 15H4-expressing construct DNA were stabilized in SFM or SPF, facilitating antibody purification for clinical use.

The epitope recognized by mAbs 2P24 and 15H4 to HCV HVR1 is located at the C terminus of HVR1 and based on the G--Q motif (positions 23–26). This was confirmed by the fact that mAb recognition of HVR1 no longer occurred after substitution of G at position 23 or Q at position 26 with V or L, respectively (Li et al., 2001). Four 9mer HVR1 peptides spanning positions 19–27 competitively inhibited the 15mer HVR1 peptide MH2 binding to cAb 2P24, but not to cAb 15H4 (Fig. 1). The results suggested that cAb 2P24 essentially reacts with the conserved G--Q epitope of HVR1. In contrast, cAb 15H4 interacts with a broader region of the C terminus of HVR1, as determined previously with the parental mAbs.

The affinity of cAbs and mAbs was generally similar, except that cAb 15H4 did not react with S85 HVR1 peptide, whereas mAb 15H4 did. This feature was retained in the cAb recognition of HVR1. A mixture of the two biotinylated cAbs or mouse mAbs reacted almost equally with a panel of 37 HVR1 peptides. The slight discrepancy in S/CO values in some HVR1 peptide EIAs between cAbs and mAbs was seen in both directions and could be related to slightly different affinities between cAbs and mAbs.

The capacity of cAbs to cross-react broadly with wild-type HCV variants was demonstrated by HCV capture. Eighty-nine per cent of HCV strains were captured by cAbs, which was consistent with the 86 % of HVR1 peptides recognized by cAbs, suggesting that the recognition of cAbs for HCV is based on the conserved HVR1 epitope exposed on HCV. However, some HCV strains containing PGAKQN in the HVR1 sequence, such as UKEH, G4720 and US (Li et al., 2001; Yanagi et al., 1997), were captured at low levels (ranging from 0·9 to 4·8 % in Table 3) by cAbs, but their corresponding HVR1 peptides containing the same sequence reacted strongly with cAbs, indicating that some particular HVR1 peptides might not fully represent the partially conformational epitopes of HVR1 on wild-type HCV particles. The percentages of captured HCV from IgG-depleted HCV plasmas were much higher than from native plasmas, suggesting that mostly uncomplexed (free) HCV was captured. Free HCV is considered the infectious portion of HCV. HCV genotypes 1, 2 and 3 were used in the capture assay and the results showed no difference among genotypes, suggesting that the capture was independent of HCV genotype.

The lack of HCV replication in cell culture limits the measurement of protective antibodies to HCV by classical neutralization assays. In our study, we used an alternative blocking assay for predicting the potentially protective ability of antibodies to HCV. The ability of cAbs to inhibit HCV binding to human target cells in vitro was measured by native and IgG-depleted HCV plasmas. At low concentrations (0·8 µg ml–1), cAbs blocked HCV binding to Molt-4 cells substantially. This function, associated with a high capacity to capture HCV, suggests that cAbs to HVR1 possess a strong potential to neutralize HCV.

One of several strategies for immunotherapy or prophylaxis of HCV infection is to prevent the binding of infectious virus to target cells. Neutralizing mAbs with a broad cross-reactivity to HCV are suitable for that purpose (Mondelli et al., 2003). The blocking and broadly cross-reactive cAbs to HVR1 presented here might be of substantial help in preventing nosocomial infection and transplanted liver reinfection by HCV.


   ACKNOWLEDGEMENTS
 
The authors thank Drs T. Grunwald and I. Harmer for providing the primers for isolation of variable-region fragments of IgG, Dr K. Armour for providing the vectors for expressing chimeric antibodies, Dr H. J. Alter for providing HCV-infected chimpanzee plasmas and Dr D. Candotti for providing the reagents used in quantitative RT-PCR. This work was supported in part by grant BS 01/4/RB18 from the National Blood Service, UK.


   REFERENCES
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
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Received 21 January 2005; accepted 17 February 2005.



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