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Pradimicin S (PRM-S) is a highly water-soluble, negatively charged derivative of the antibiotic pradimicin A (PRM-A) in which the terminal xylose moiety has been replaced by 3-sulfated glucose. PRM-S does not prevent human immunodeficiency virus (HIV) adsorption on CD4+ T cells, but it blocks virus entry into its target cells. It inhibits a wide variety of HIV-1 laboratory strains and clinical isolates, HIV-2, and simian immunodeficiency virus (SIV) in various cell culture systems (50% and 90% effective concentrations [EC50s and EC90s] invariably in the lower micromolar range). PRM-S inhibits syncytium formation between persistently HIV-1- and SIV-infected cells and uninfected CD4+ T lymphocytes, and prevents dendritic cell-specific intercellular adhesion molecule-3-grabbing nonintegrin (DC-SIGN)-mediated HIV-1 and SIV capture and subsequent virus transmission to CD4+ T cells. Surface plasmon resonance (SPR) studies revealed that PRM-S strongly binds to gp120 in a Ca2+-dependent manner at an affinity constant (KD) in the higher nanomolar range. Its anti-HIV activity and HIV-1 gp120-binding properties can be dose-dependently reversed in the presence of an (α-1,2)mannose trimer. Dose-escalating exposure of HIV-1-infected cells to PRM-S eventually led to the isolation of mutant virus strains that had various deleted N-glycosylation sites in the envelope gp120 with a strong preference for the deletion of the high-mannose-type glycans. Genotypic resistance development occurred slowly, and significant phenotypic resistance occurred only after the sequential appearance of up to six mutations in gp120, pointing to a high genetic barrier of PRM-S. The antibiotic is nontoxic against a variety of cell lines, is not mitogenic, and does not induce cytokines and chemokines in peripheral blood mononuclear cells as determined by the Bio-Plex human cytokine 27-plex assay. It proved stable at high temperature and low pH. Therefore, PRM-S may qualify as a potential anti-HIV drug candidate for further (pre)clinical studies, including its microbicidal use.
Carbohydrate-binding agents (CBAs) comprise a broad and structurally diverse functional class of agents. The most abundant representatives among the CBAs are the lectins (2). The innate immune system uses lectins for capturing and processing pathogen antigens to destroy the pathogen and/or to trigger a proper immune response against the pathogen (i.e., dendritic cell-specific intercellular adhesion molecule-3-grabbing nonintegrin [DC-SIGN], langerin, mannose-binding lectin [MBL], macrophage mannose receptor [MMR], defensins) (9, 11-13, 32). Interestingly, a variety of lectins derived from several organisms different from mammalians have been described to be endowed with anti-HIV activity. Such CBAs include plant (i.e., Hyppeastrum hybrid agglutinin [HHA], Galanthus nivalis agglutinin [GNA], and Urtica dioica agglutinin [UDA]), invertebrate (i.e., mermaid lectin), and prokaryotic (i.e., cyanovirin N, actinohivin, cytovirin) lectins (for an overview see reference 2). These compounds have been shown to inhibit infection of cells by cell-free virus and to prevent syncytium formation in cocultures of HIV-infected and uninfected T lymphocytes, DC-SIGN- or MMR-directed virus capture, and subsequent virus transmission to T lymphocytes (7, 25). In this respect, the CBAs may qualify as potential anti-HIV drugs from both a systemic and a microbicidal viewpoint. However, the above-mentioned CBAs are proteins whose molecular masses range between 8,700 Da (i.e., UDA) and 50,000 Da (i.e., HHA, GNA) or even higher. Such proteins may have a variety of disadvantages to become potential drugs, including efficient scale-up and purification of bulk amounts, poor (if any) oral bioavailability, susceptibility to proteases, short plasma half-life, and potential generation of an immune response.
Two decades ago, nonpeptidic, low-molecular-weight antibiotics designated pradimicin A (PRM-A) and benanomycin A (BNM-A) were discovered in the culture fluid of Actinomadura hibisca (24) and Actinomadura sp. MH 193-16F4 (16, 20). PRM-A inhibited the growth of fungi (such as Aspergillus) (29) and behaved like the above-mentioned lectins: efficient inhibition of HIV infection and prevention of syncytium formation between persistently HIV-1-infected and uninfected cells (28). Interestingly, we recently could demonstrate that PRM-A, similar to lectins, selects for virus mutants in cell cultures containing deletions in N-glycosylation sites of gp120 (8). Thus, the nonpeptidic PRM-A antibiotic acts as a lectin in terms of glycan recognition, antiviral activity, and drug resistance pattern. Since PRM-A is endowed with a limited solubility (≤50 μM), we searched for more soluble PRM-A analogues and we focused our research efforts on a modified PRM-A derivative, produced by a related microorganism (Actinomadura spinosa strain A A08 51) that contains a negatively charged 3-sulfated glucose as its terminal sugar instead of the uncharged xylose in PRM-A (26, 27) (Fig. (Fig.11).
We investigated the mechanism of action and the antiviral properties of PRM-S in various cell culture systems. Based on our findings, we propose this compound to be a most suitable drug candidate for further studies with monkeys that are intravaginally or systemically challenged by SIV to evaluate for the first time a prototype nonpeptidic small-size CBA in the in vivo setting both as a potential microbicide and for systemic administration.
The pradimicin strains Actinomadura sp. TP-A0019 and Actinomadura sp. TP-A0020 were isolated from soil collected in Toyama, Japan. Pradimicin A (PRM-A) and pradimicin S (PRM-S) (Fig. (Fig.1),1), respectively, were produced by these strains and isolated and purified from the culture supernatants as previously described (24, 26). Hyppeastrum hybrid agglutinin (HHA), Galanthus nivalis agglutinin (GNA), and Urtica dioica agglutinin (UDA) were kindly provided by E. Van Damme (Ghent, Belgium). Tenofovir was provided by A. Holý (Prague, Czech Republic). The bicyclam AMD3100 was purchased from Sigma (St. Louis, MO).
Human T-lymphocytic C8166, Molt4/C8, and CEM cells and cervical carcinoma HeLa, colon carcinoma CaCo-2, and glioblastoma U87 cells were obtained from the American Type Culture Collection (Manassas, VA). MT-4 cells were provided by L. Montagnier (at that time at the Pasteur Institute, Paris, France). The Raji/DC-SIGN cells were constructed by Geijtenbeek et al. (14). Wild-type Raji/0 and DC-SIGN-expressing Raji/DC-SIGN cells were kindly provided by L. Burleigh (Institut Pasteur, Paris, France). All cell lines mentioned were cultivated in RPMI 1640 medium supplemented with 10% fetal bovine serum (FBS) (BioWhittaker Europe, Verviers, Belgium), 2 mM l-glutamine, and 0.075 M NaHCO3.
HIV-1 (strains IIIB and BaL) was provided by R. C. Gallo and M. Popovic (Institute of Human Virology, University of Maryland, Baltimore, MD). HIV-2 (ROD) was provided by L. Montagnier (at that time at the Pasteur Institute, Paris, France). SIVmac251 was obtained from C. Bruck. The primary clinical isolates representing different HIV-1 clades and an HIV-2 isolate were all kindly provided by L. Lathey from BBI Biotech Research Laboratories, Inc., Gaithersburg, MD, and their coreceptor use (R5 or X4) was determined.
The human T4 lymphocytic C8166, Molt4/C8 and CEM cells and cervical carcinoma HeLa, colon carcinoma CaCo-2, and glioblastoma U87 cells were seeded in 96-well (200-μl) microtiter plates at 20,000 to 75,000 cells per well in the presence of different concentrations of PRM-S and PRM-A. After 3 to 4 days of incubation at 37°C, the cells were counted in a Coulter Z1 particle counter (Analis, Ghent, Belgium). The 50% cytostatic concentration (CC50) was defined as the compound concentration required to inhibit tumor cell proliferation by 50%.
CEM and C8166 cells (5 × 105 cells per ml) were suspended in fresh culture medium and infected with HIV-1 at 100 50% cell culture infective doses (CCID50) per ml of cell suspension. Then, 100 μl of the infected cell suspension was transferred to microplate wells, mixed with 100 μl of the appropriate dilutions of the test compounds, and further incubated at 37°C. After 3 to 4 days, giant cell formation was recorded microscopically in the cell cultures and the number of giant cells was estimated as a percentage of the number of giant cells present in the nontreated virus-infected cell cultures (~50 to 100 giant cells in one microscopic field when examined at a microscopic magnitude of ×100). The 50% effective concentration (EC50) corresponds to the compound concentrations required to prevent syncytium formation in the virus-infected C8166 and CEM cell cultures by 50%.
MT-4 cells (5 × 105 cells/ml) were suspended in fresh culture medium and infected with SIV at 100 CCID50 per ml cell suspension (1 CCID50 being the dose infective for 50% of the cell cultures). Then, 100 μl infected cell suspension was transferred to microtiter plate wells, mixed with 100 μl of the appropriate dilutions of compound, and further incubated at 37°C. The number of viable cells was determined by trypan blue staining for both virus-infected and mock-infected cell cultures using a KOVA Glasstic Slide 10 with a quantitative grid (Hycor Biomedical Ltd., Penicuik, United Kingdom). The 50% effective concentration (EC50) and 50% cytostatic concentration (CC50) were defined as the compound concentrations required to reduce the number of viable cells in the virus-infected and mock-infected cell cultures, respectively, by 50%.
PBMCs from healthy donors were stimulated with PHA at 2 μg/ml (Sigma, Bornem, Belgium) for 3 days at 37°C. The PHA-stimulated blasts were then seeded at 0.5 × 106 cells per well into a 48-well plate containing various concentrations of compound in cell culture medium (RPMI 1640) containing 10% fetal calf serum (FCS) and interleukin-2 (IL-2) (25 U/ml; R&D Systems Europe, Abingdon, United Kingdom). The virus stocks were added at a final dose of 250 pg p24 or p27/ml. Cell supernatant was collected at day 12 and HIV-1 core antigen (Ag) in the culture supernatant was analyzed by a p24 Ag enzyme-linked immunosorbent assay (ELISA) kit (Perkin Elmer, Zaventem, Belgium). For HIV-2 p27 Ag detection, the Innotest from Innogenetics (Temse, Belgium) was used. Mock-infected cell cultures exposed to the different drug concentrations were examined for cell viability by trypan blue staining at day 7 of incubation.
MT-4 cells or Raji/DC-SIGN cells (5 × 106) were incubated with compound (20 μM) or medium for 30 min at 4°C. Next, supernatant containing 1 × 105 pg of p24 Ag of HIV-1(NL4.3) virus was added. One hour later, cells were washed three times with medium and lysed, and p24 Ag bound to the cells was determined by a specific p24 Ag ELISA.
Persistently SIVmac251- or HIV-1 (IIIB)-infected HUT-78 cells (designated HUT-78/SIV or HUT-78/HIV-1) were generated by infection of 7-ml cultures of 1 × 105 HUT78 cells/ml with SIVmac251 (~800,000 pg) or HIV-1 (IIIB) (~800,000 pg). The cells were subcultured every 3 to 4 days, and persistent virus infection was monitored in the culture supernatants using the SIV p27 or HIV p24 antigen enzyme-linked immunosorbent assay kit (Gentaur, Brussels, Belgium).
For the cocultivation assays, HUT-78/SIV or HUT-78/HIV-1 cells were washed to remove free virus from the culture medium, and 5 × 104 cells (50 μl) were transferred to 96-well microtiter plates. Then, a similar amount of C8166 cells (50 μl), along with an appropriate concentration of test compound (100 μl), was added to each well. After 2 days, the EC50s were determined based on the appearance of syncytia in the cocultures.
Exponentially growing B-lymphocyte Raji wild-type (Raji/0) and DC-SIGN-expressing (Raji/DC-SIGN) cells were suspended in cell culture medium at 6 × 106 cells/400 μl as described by Balzarini et al. (7). Then, 0.4-ml cell suspensions were exposed to 600 μl wild-type HIV-1 (IIIB) (2.2 × 106 pg/ml p24) for 60 min, after which 39 ml culture medium was added to the virus-infected cell culture. The cells were centrifuged at 1,250 rpm for 10 min, 39.9 ml supernatant was carefully removed, and the virus-exposed cells were resuspended in 40 ml medium. After a second centrifugation step, 39.9 ml supernatant was again removed, and the remaining 0.1 ml cell suspension was 10-fold diluted in cell culture medium to 1 ml. Under these experimental (washing) conditions, a maximum of 8 pg HIV-1 p24 could have remained in the 1-ml supernatant (or 0.4 pg in 50 μl). A 50-μl cell suspension was withdrawn for p24 Ag determination by a specific HIV-1 p24 ELISA, and 50 μl of the Raji/DC-SIGN cell suspension was added to 96-well microplates in which 100-μl compound dilutions were present. Then, 50 μl C8166 cells (107/ml) was added to each well. These mixed cell cultures were incubated at 37°C in a CO2-controlled humidified incubator and microscopically scored for syncytium formation at ~36 to 48 h post-virus exposure/cocultivation. It should be mentioned that the maximum amount of virus that could have remained in the culture medium (<1 pg HIV-1 p24) is unable to result in HIV-1-induced giant cell formation in C8166 cell cultures within the time period of analysis (36 to 48 h).
Large amounts of HIV-1 particles [100 μl; ~2.2 × 106 pg p24/ml X4 HIV-1 (IIIB) or R5 HIV-1 (BaL)] or SIV particles (100 μl; ~1.5 × 105 pg p27/ml) were exposed to serial dilutions of the test compounds (400 μl) for 30 min as described before (7) Then, the drug (i.e., PRM-A, PRM-S, and as a positive control HHA )-exposed virus suspensions (500 μl) were mixed with Raji/DC-SIGN cell suspensions (500 μl; 106 cells) for 60 min at 37°C, after which the cells were thoroughly washed twice with 40 ml culture medium as described above. This procedure resulted in a final dilution of the initial compound concentrations by at least 160,000-fold. The Raji/DC-SIGN cell cultures were then analyzed for p24 Ag content. Also, 200 μl of the cell cultures was mixed with 2 × 105 C8166 cells (800 μl) (resulting in a further 5-fold dilution of the original amount of exposed test compound) and further incubated in 48-well plates for 36 to 48 h at 37°C. Then, the syncytium formation in the cell cultures was evaluated microscopically.
Recombinant gp120 protein from HIV-1 strain IIIB (ImmunoDiagnostics Inc., Woburn, MA) (produced by CHO cell cultures) and recombinant gp41 protein from HIV-1 (HXB2 strain) (Acris Antibodies GmbH, Herford, Germany) (produced by Pichia pastoris) were covalently immobilized on the carboxymethylated dextran matrix of a CM5 sensor chip in 10 mM sodium acetate, pH 4.0, using standard amine coupling chemistry to final densities of 122 RU (~1 fmol of gp120) and 58 RU (~1.4 fmol), respectively. A reference flow cell was used as a control for nonspecific binding and refractive index changes. All interaction studies were performed at 25°C on a Biacore T100 instrument (GE Healthcare, Uppsala, Sweden). The compounds PRM-A and PRM-S were serially diluted in HBS-P (10 mM HEPES, 150 mM NaCl, and 0.05% surfactant P20; pH 7.4), supplemented with 5% dimethyl sulfoxide (DMSO, Merck) for PRM-A only, and with 10 mM CaCl2 covering a wide concentration range (2.5 to 40 μM) by using 2-fold dilution steps. Samples were injected for 2 min at a flow rate of 45 μl/min, and the dissociation was monitored for 6 min. Several buffer blanks were used for double referencing. A DMSO concentration series was included to eliminate the contribution of DMSO to the measured response. The CM5 sensor chip surface was regenerated with one injection of 50 mM NaOH. All studied interactions (PRM-A, PRM-S) on gp120 IIIB and gp41 HXB2 result in a specific binding signal. The shape of the association and dissociation phases reveals that the curves are not according to a perfect 1:1 Langmuir adsorption model. The experimental data were fit using the 1:1 binding model (Biacore T100 Evaluation software, verison 2.0.1) to determine the binding kinetics. These affinity and kinetic values are apparent values, as the injected concentrations of the evaluated compounds did not result in complete saturation of the binding signals and the dissociation phases are biphasic (see Results).
In another set of experiments, HIV-1 gp120 was coupled to a CM5 sensor chip up to an immobilization level of 457 response units (RU) (3.8 fmol of gp120) as described above. The experimental setup of the inhibition studies was identical to the method described for the kinetic experiments. PRM-S was diluted in HBS-P buffer supplemented with 10 mM CaCl2. The compound, alone or in the presence of (α-1,2)mannose trimer, was injected for 3 min over the chip at a flow rate of 30 μl/min, after which the compound was allowed to dissociate spontaneously for 5 min in the presence of HBS-P buffer supplemented with 10 mM CaCl2. The inhibition of the binding of PRM-S to gp120 by (α-1,2)mannose trimer was tested with a wide range of trisaccharide concentrations. An injection of (α-1,2)mannose trimer at a concentration of 200 μM (in the absence of PRM-S) was included as a control.
In three independent experiments, PBMCs and PHA-blasts (derived from three different donors) were cultured in the presence of 22 and 21 μM PRM-A and PRM-S, respectively, and culture supernatants were collected after 72 h. The cytokine production profile was determined by the Bio-Plex 200 system (Bio-Rad, Hercules, CA) and Bio-Plex human cytokine 27-plex assay according to the manufacturer's instructions and also as described by Huskens et al. (19). The 27-plex assay kit contains beads conjugated with monoclonal antibodies (MAbs) specific for interleukin-1α (IL-1α), IL-1ra, IL-2, IL-4, IL-5, IL-6, IL-7, IL-8, IL-9, IL-10, IL-12, IL-13, IL-15, IL-17, eotaxin, fibroblast growth factor (FGF), granulocyte colony-stimulating factor (G-CSF), granulocyte-macrophage-CSF (GM-CSF), gamma interferon (IFN-γ), interferon-inducible protein-10 (IP-10), monocyte chemoattractant protein-1 (MCP-1), macrophage inflammatory protein-1α (MIP-α), MIP-1β, platelet-derived growth factor-BB (PDGF-BB), regulated on activation normal T-cell expressed and secreted (RANTES), tumor necrosis factor alpha (TNF-α), and vascular endothelial growth factor (VEGF). For each cytokine, nine standards ranging from approximately 0.5 pg/ml to 32,000 pg/ml were constructed and the minimum detectable dose was between 0.5 pg/ml and 5 pg/ml. Standard curves and the concentration of cytokines within samples were generated with the Bio-Plex Manager 4.1 software.
HIV-1 (IIIB) was added to CEM cell cultures in 48-well plates in the presence of PRM-S at a concentration equal to 1 to 2 times the EC50. For the generation of drug-resistant virus mutants, an increased drug concentration was administered when full cytopathicity was obtained in the previous cell culture as described before (8). The drug resistance selection schedule is depicted in Fig. Fig.2.2. Virus isolates from subculture passages 14 and 42 were taken during the drug resistance selection process and further genotypically and phenotypically characterized.
Proviral DNA was extracted from cell pellets using the QIAamp blood minikit (Qiagen, Hilden, Germany). Genotyping of both the gp120 and gp41 genes was determined in this assay as described before (33).
PRM-S was dissolved in phosphate-buffered saline (PBS) at 1 mM and incubated at 50°C for 4 days or was dissolved in 7 mM sodium phosphate buffer, pH 4.0, and kept at room temperature for 4 days. After the incubation period, the compound solution was analyzed for stability by high-pressure liquid chromatography (HPLC) analysis on a Merck (Darmstadt, Germany) LiChroCART 125-4 RP column (5 μm) using the following gradients (flow, 1 ml/min): 2 min at 98% buffer (50 mM NaH2PO4 [Acros, NJ] plus 5 mM heptanesulfonic acid [Sigma, St. Louis, MO], pH 3.2) and 2% acetonitrile (ACN) (Biosolve, Valkenswaard, Netherlands); 6 min linear gradient to 80% buffer and 20% ACN; 2 min linear gradient to 75% buffer and 25% ACN; 10 min linear gradient to 65% buffer and 35% ACN; 10 min linear gradient to 50% buffer and 50% ACN; 10 min isocratic flow; 5 min linear gradient to 98% buffer and 2% ACN; and 5 min equilibration under the same conditions. The retention time of PRM-S was 16.8 min.
PRM-S inhibited the cytopathic effect of HIV-1 (IIIB) and HIV-2 (ROD) in a variety of laboratory CD4+ cell lines, including MT-4, CEM, and C8166 cells (Table (Table1).1). Its 50% effective concentrations (EC50) ranged between 5.1 and 8.9 μM. Under comparable experimental conditions, the EC50 of PRM-A ranged between 5.2 and 5.9 μM (Table (Table1).1). Both PRM-S and PRM-A also consistently suppressed a wide variety of CCR5-tropic (R5) and CXCR4-tropic (X4) HIV-1 clade isolates, as well as HIV-2 in peripheral blood mononuclear cell (PBMC) cultures at EC50s that ranked in the lower micromolar range (3.1 to 18 μM for PRM-S and 1.6 to 10 μM for PRM-A) (Table (Table2).2). Their EC90 values were only 2- to 3-fold higher than their EC50s (data not shown), suggesting a relatively steep dose-response curve. PRM-S and PRM-A also inhibited SIVmac251 replication in MT-4 cell cultures at EC50s of 8.1 and 5.0 μM, respectively. The antibiotics were not cytostatic against a variety of different cell lines, such as murine leukemia L1210; human lymphocyte CEM, C8166, and Molt4/C8; cervical carcinoma HeLa; osteosarcoma OST/TK−; colon carcinoma CaCo-2; and glioblastoma U87 cells and also primary PBMCs (CC50s varied depending on the nature of the cell lines and ranged for PRM-S between 230 and >500 μM; for PRM-A, CC50s were invariably >50 μM [solubility limit]). Viability of the drug-exposed PBMCs at day 7 of incubation was >95% at 500 μM PRM-S and 50 μM PRM-A.
PRM-S and PRM-A also efficiently prevented syncytium formation between persistently HIV-1 (IIIB)- or SIVmac251-infected HUT-78 cells and uninfected SupT1 or CEM cells (Table (Table3).3). Both drugs showed EC50s ranging between 3.4 and 12 μM, that is, at concentrations that also prevented the infection of T-lymphocyte cells by cell-free virus particles.
The potential of PRM-S and PRM-A to prevent capture of X4 HIV-1 (IIIB) and R5 HIV-1 (BaL) by DC-SIGN has been investigated using Raji cells that were transfected with the DC-SIGN gene (14, 15) and abundantly express DC-SIGN in their cell membranes (14, 15). Short exposure of Raji/DC-SIGN cell cultures to HIV-1 (IIIB) significantly captured the virus to their cell membranes (7). Nontransfected wild-type Raji/0 cells did not measurably capture HIV-1. Different pradimicin concentrations were shortly (30 min) preexposed to HIV-1 before the virus was administered to the Raji/DC-SIGN cells. After 60 min, unadsorbed virus and test compound were carefully removed by serial washing steps and the amount of captured virus was determined by measurement of the p24 content of the cells. Both PRM-S and PRM-A efficiently and dose-dependently inhibited binding of HIV-1 to the DC-SIGN cells. Their 50% inhibitory concentrations (IC50s) were 11 μM and 3.5 μM for the X4 HIV-1 (IIIB) and 21 μM and 17 μM for the R5 HIV-1 (BaL) strains, respectively. The α(1-3)/α(1-6)-specific plant lectin HHA was included as a positive control and found to be 20- to ~100-fold more effective (Table (Table44).
When the washed, drug-treated, virus-exposed DC-SIGN+ cells were cocultured with uninfected C8166 cells, abundant syncytium formation occurred within 24 and 48 h postcocultivation when the captured virus had not been preexposed to the drugs. In contrast, in the cocultures in which the virus had been preexposed to different drug concentrations before being captured by Raji/DC-SIGN cells, both PRM-S and PRM-A dose-dependently inhibited syncytium formation at EC50s of 7.1 and 3.4 μM, respectively. HHA was 5- to 10-fold more effective (Table (Table44).
In another set of experiments, virus (HIV-1 strain IIIB) was given the opportunity to be captured by Raji/DC-SIGN cells. Then, the HIV-1-captured Raji/DC-SIGN cells were cocultured with C8166 cells in the presence of a variety of test compound concentrations. Giant cell formation (appearing upon transmission of HIV-1 from the Raji/DC-SIGN to the C8166 cells) was recorded as a parameter of efficiency of virus transmission. The PRM-S and PRM-A derivatives efficiently prevented transmission of virus captured by DC-SIGN-expressing Raji cells to T-lymphocyte C8166 cell cultures (IC50, 5.1 to 5.4 μM) (Table (Table4).4). In conclusion, PRM-S and PRM-A were able to efficiently prevent virus capture by DC-SIGN-expressing cells and subsequent transmission to, and infection of, T lymphocytes. Comparable data were obtained when the inhibitory effect of the antibiotics on SIVmac251 capture by Raji/DC-SIGN and subsequent virus transmission to C8166 cells was determined (IC50, 16 ± 7.4 μM).
PRM-S was investigated for its binding potential against HIV-1 gp120 and gp41 and compared with PRM-A using surface plasmon resonance (SPR) technology. For this purpose, HIV-1 (IIIB) gp120 and HIV-1 (HXB2) gp41 were covalently immobilized on the sensor chip, after which the envelope glycoproteins were exposed to serial dilutions of PRM-S or PRM-A (range, 2.5 μM to 40 μM). In the absence of CaCl2, the antibiotics did not show any binding capacity to gp120 or gp41 (data not shown). However, in the presence of 10 mM CaCl2, both PRM-S and PRM-A showed comparable binding capacities to gp120 (amplitude, ~90 to 110 RU at 40 μM after 120 s) (Fig. (Fig.3A3A).
The binding of the pradimicins to gp120 clearly showed biphasic dissociation kinetics characterized by an initial fast dissociation rate followed by a much slower dissociation at later time points (>30 s after the start of dissociation) (Fig. (Fig.3A).3A). The apparent kd (dissociation rate constant) rate values (and KD and ka [association rate constant]) were calculated from the three lowest drug concentrations covering a 360-s time period after replacement of the pradimicins in the solute by buffer, using a 1-to-1-binding model. The affinity constant KD was identical for PRM-S and PRM-A (KD, ~0.40 μM), as also observed for the kd and ka values. Binding of PRM-S to HIV-1 gp120 could be dose-dependently reversed by an (α-1,2)mannose trimer (Fig. (Fig.3B).3B). When similar kinetic determinations were performed for gp41, 30- to 35-fold higher apparent KD values were found than for gp120. This was due to an ~5-fold higher kd but a 10- to 20-fold lower ka for the gp41 than for the gp120 interaction (Fig. (Fig.3A;3A; Table Table55).
HIV-1 (IIIB)-infected CEM cell cultures were exposed to PRM-S at 10 μM, representing a 1 to 2 times the EC50 of PRM-S. Every 4 to 5 days, subcultivations were performed. The PRM-S concentration was stepwise increased only after abundant syncytium formation occurred (Fig. (Fig.2).2). At two time points during the selection process (i.e., passage 14 [~63 days] [at 10 μM PRM-S] and passage 42 [~189 days] [at 100 μM PRM-S]), virus isolates were taken for genotypic and phenotypic characterization. The virus isolates contained mutations in N-glycosylation motifs of HIV-1 gp120 but not of gp41. The number of mutations in the viral envelope was higher at the longer (drug-escalating) selection time. The virus isolate at passage 14 contained two N-glycan deletions in gp120, whereas the virus isolate at the end of the selection (passage 42) contained up to six different deletions in N-glycosylation motifs (Table (Table6)6) . In the virus isolate (number 14), a new glycosylation site created by a 5-amino acid insertion appeared, but this site was not present anymore in the later isolate (number 42) (Table (Table6).6). In all cases (except the N392D mutation) the S or T of the glycosylation motif had been replaced by another amino acid, thereby destroying the glycosylation motif. The mutated glycosylation sites with deleted glycans (shown as red balls in Fig. Fig.4)4) were spotted on the three-dimensional structure of HIV-1 gp120 (amino acid numbering as described by Kwong et al. ). It became evident that in the mutant HIV-1 strains (i) none of the six (complex-type) glycans in the V1/V2 area of gp120 were deleted and (ii) there was a striking preference for the deletion of high-mannose-type glycans in the remaining envelope part. Indeed, except for the N-88 mutation, all other glycosylation site mutations exclusively affected high-mannose-type glycans (as originally determined by Leonard et al. ) (Fig. (Fig.4;4; Table Table6).6). It should be noticed that, beside the replacement of the N or T/S amino acids in the glycosylation motifs, additional mutations at nonglycosylation sites were observed in both virus isolates, appearing mostly as mixtures with wild-type virus (i.e., S164N/S and V208I/V in isolate number 14 and K135K/E, S164N/S, N340N/Y, N302Y, and K337E in isolate number 42). No glycosylation site mutations were recorded in gp41 of the virus isolates.
Sensitivity of the mutant HIV-1 isolates to the inhibitory effect of PRM-S and its closely related PRM-A derivative was determined (Table (Table7).7). Whereas the virus isolate (number 14) barely lost its sensitivity for the inhibitory activity of the pradimicins, the virus isolate (number 42) that contained six glycan mutations showed a ≥20-fold phenotypic resistance to the pradimicins (EC50 > 50 μM). The PRM-S-resistant virus strain proved also phenotypically cross-resistant to other CBAs, such as the entry inhibitors HHA, GNA, and UDA, but not to the CXCR4 coreceptor antagonist AMD3100 (Table (Table77).
The infectivity and replication potential of the HIV-1 mutant isolate number 42 have also been determined by exposing MT-4 cell cultures to p24 amounts equal to those used for HIV-1/WT and HIV-1/PRM-S number 42 and measuring viral p24 production in the supernatants of the infected cell cultures as a function of time. The virus production in wild-type virus-infected cells proceeded much earlier and faster than in the mutant virus-infected cells (Fig. (Fig.55).
PRM-S and PRM-A were investigated for their potential to induce chemokines and cytokines in PBMCs. For this purpose, the Bio-Plex system, which quantifies simultaneously a wide variety of different cytokines (IL-1β, IL-1ra, IL-2, IL-4, IL-6, IL-8, IL-9, IL-10, IL-12, IL-13, IL-15, eotoxin, FGF, G-CSF, GM-CSF, IP-10, MCP-1, MIP-1α, MIP-1β, PDGF, RANTES, TNF-α, and VEGF) in one single sample, was used (10). Both PRM-S and PRM-A were used at 20 μg/ml (21 and 22 μM, respectively), that is, at drug concentrations that exceed the EC50 against HIV-1 by 3- to 10-fold. Neither pradimicin antibiotic seemed to have any stimulatory effect on the chemo/cytokine production levels in PBMC (data not shown). Instead, there was rather a slight trend to suppress cytokine production at 72 h post-drug administration (~4-fold decrease for IL-2 and RANTES and ~2- to 3-fold decrease for IL-4, IL-9, IL-13, IL-15, eotoxin, G-CSF, and PDGF). In contrast, the prokaryotic (α-1,2)mannose-specific lectin cyanovirin N (CV-N) was previously shown to dramatically stimulate the production of a wide variety of cytokines and chemokines in PBMCs at 0.2 μM, under identical experimental conditions using the same assay system (19).
In view of the potential of PRM-S to be used as a microbicide drug lead, the stability of the compound was investigated. PRM-S (1 mM) was heated at 50°C in a PBS solution or exposed to pH 4.0 in 7 mM sodium phosphate buffer for 4 days. Then, the drug solution was analyzed by HPLC on a reverse-phase column and found to be entirely stable under these experimental conditions. Only intact compound could be detected in the samples (Rf, 16.9 min). The drug solutions were then also evaluated for their inhibitory activity against syncytium formation in HUT-78/HIV-1 and Sup T-1 cocultures. The antiviral efficacy of the compound exposed to high temperature or low pH for four subsequent days was comparable with the antiviral activity of unexposed compound (data not shown). Also, SPR analyses revealed that PRM-S still showed extensive binding to HIV-1 gp120, bound at the sensor chip, at pH 4 (B. Hoorelbeke, unpublished observation).
The stability of PRM-S and the preservation of its antiviral potential upon exposure to high temperature (4 days, 50°C) and low pH (4 days, pH 4.0) further add to its microbicidal potential. In contrast to PRM-A, which is endowed with a limited solubility in aqueous media (i.e., ~50 μM), PRM-S is an acidic antibiotic that is readily soluble in aqueous media (>20 mM at physiological pH) (27). This increased solubility of PRM-S may give the drug a therapeutic edge over the prototype PRM-A derivative in terms of formulation, attainable drug peak levels in plasma as well as locally (i.e., intravaginally), and more durable suppression of virus infection at concentrations higher than 50 μM, a property which may be of crucial importance from a microbicidal viewpoint of drug application. However, since PRM-S is an antibiotic that recognizes glycans but was found not to show any effect on the growth of Staphylococcus aureus, Bacillus subtilis, or Escherichia coli (26), it would be important to investigate its effect on the normal flora of the cervicovaginal tract, including Lactobacillus spp., before further development as a potential microbicidal agent. Although the antiviral activity of PRM-S is in the lower micromolar range and less pronounced than several other drugs currently used as therapeutics or investigated as potential microbicides, it should be noticed that this type of drug has an activity range comparable to that of tenofovir, which is currently included in microbicide trials and shows to be a cornerstone drug for systemic HIV treatment. Also, PRM-S does not need further metabolic conversion before it becomes active, and it acts at the outside of the cells and does not need cellular uptake and metabolism before it may inhibit its target.
Our antiviral data revealed that PRM-S behaves quite similarly to the prototype PRM-A derivative in terms of its potential to inhibit (i) cell-free virus infection in cell lines and a broad range of HIV clades in PBMC, (ii) syncytium formation between persistently HIV/SIV-infected cells and uninfected cells, (iii) DC-SIGN-directed HIV/SIV capture, and (iv) subsequent virus transmission to uninfected CD4+ T lymphocytes. The latter properties are especially beneficial if the drug would be used as a microbicide, since DC-SIGN-directed entry and transmission to T lymphocytes are considered an important avenue of primary infection of individuals exposed to HIV through sexual intercourse. In this study, inhibition of virus capture by DC-SIGN has been demonstrated for an HIV-1 X4 virus strain (IIIB) and an HIV-1 R5 virus strain (BaL). It was not surprising that PRM-S also prevented HIV-1 R5 virus capture by DC-SIGN, since it was previously demonstrated that CBAs, including PRM-A, were also able to efficiently and dose-dependently prevent capture of HIV-1 R5 (BaL strain) by the macrophage mannose receptor (MMR) present on primary macrophage cells (25). It should be mentioned that, as also previously observed for other CBAs, such as several plant lectins (3), PRM-S and PRM-A do not inhibit HIV binding to the CD4 receptor (data not shown). These compounds clearly act against the virus fusion process at a stage after initial interaction of the virus to the target cells had occurred. It is, however, also important to realize that whereas the CBAs do not prevent adsorption of the virus to the CD4+ target cells, they efficiently inhibit fusion to the target cells and also binding to the DC-SIGN receptor and subsequent virus transmission to T lymphocytes.
Like PRM-A, PRM-S selects for mutant drug-resistant virus strains that contain deletions in a variety of N-glycosylation sites of HIV gp120. In fact, all glycan deletions observed in the different PRM-S-exposed HIV-1 strains have also been observed in PRM-A-exposed virus strains except the N-392 glycan deletion. In a variety of PRM-A-exposed virus isolates that had been previously analyzed for glycosylation site mutations in gp120 (8), glycan deletions at N234, N301, N332, and N448 that have not been found in the PRM-S-exposed HIV-1 strains were also observed. However, it is not clear whether these apparent differences between PRM-S and PRM-A are relevant, and it may well be possible that these differences disappear if additional mutant virus strains should be selected in the presence of escalating PRM-S concentrations. Whereas a variety of N-glycosylation sites were detected in HIV-1 gp120 under pradimicin drug pressure, no consistent glycan deletions were observed in HIV-1 gp41, although the antibiotics also bind to gp41 (as evident from our SPR studies). Traces of one glycan deletion in gp41 had been observed in a viral isolate mixture at 7 μg/ml PRM-S (not shown), but this mutation was not further retained at higher drug concentrations (i.e., in the isolates at 10 and 100 μg/ml PRM-S) (Table (Table6).6). This predominant lack of gp41 glycan deletions has also been observed for other CBAs (i.e., HHA, GNA, UDA, cyanovirin N, 2G12 [4-6, 8, 18]) and may reflect the fact that gp41 might not be available for drug interaction during the initial virus binding and entry steps. Thus, the selection pressure of other CBAs and also PRM-S acts predominantly on gp120 but not on gp41.
It is striking that five out of the six glycosylation sites that were mutated in HIV-1 gp120 under PRM-S pressure (isolate number 42) were reported to contain high-mannose-type glycans. Given the fact that 11 out of the 24 glycosylation sites in gp120 were determined to contain a high-mannose-type or hybrid-type glycan (46%) (22), it seems clear that PRM-S predominantly selects for deletions of these types of glycans and, thus, may likely show a strong preference for binding to high-mannose-type glycans. Such a preference was also earlier observed for PRM-A (8). The ability of an (α-1,2)mannose trimer to dose-dependently decrease the interaction of PRM-S with HIV-1 gp120, as demonstrated in the SPR experiments, is in full agreement with the reported binding of pradimicin to high-mannose-type glycans [containing (α-1,2)mannose residues] (28). (α-1,2)Mannose bonds are abundantly present at the terminal sugars of high-mannose-type glycans, but they are completely absent in complex-type glycans. This may mean that the carbohydrate bond preferences of the pradimicin antibiotics are similar to those observed for the prokaryotic cyanovirin-N but differ from those of the HHA and GNA plant lectins that were reported to predominantly show α(1,3) and/or α(1,6)mannose oligomer preference (31). Our observations may also explain why none of the six complex-type glycans present in the HIV-1 V1/V2 gp120 loop have been found mutated under PRM-S (or PRM-A ) pressure and why PRM-A has previously been shown to keep its full inhibitory potential against HIV-1 strains that contain a variety of glycan deletions in V1/V2 (1). Alternatively, it cannot be excluded that the pradimicins can better reach the high-mannose-type glycans that seem to be preferentially clustered at the outer domain of gp120 than the complex-type glycans that are preferentially located at the less-accessible inner domains of gp120.
As has also been previously observed for some HIV-1 strains that had emerged under CBA (i.e., HHA and GNA) pressure and in which a variety of N-glycans were deleted in gp120, and for mutant HIV-1 strains in which multiple envelope gp120 glycan deletions had been introduced in V1/V2 (1), the infectivity potential and virus production were compromised for the PRM-S-resistant virus isolate compared to those for the wild-type virus (Fig. (Fig.5).5). The forced accumulation of N-glycan deletions in HIV-1 gp120 and the resulting lower infectivity potential of such mutant virus strains by pressure of PRM-S to the virus-infected cell culture are interesting phenomena from a clinical viewpoint. These phenomena may open novel therapeutic perspectives for such drugs, since the accumulation of drug resistance mutations (i.e., N-glycan deletions) may not only result in a lower infectivity potential of the virus but also, and at least equally if not more importantly, trigger the immune system to elicit neutralizing antibodies and/or a cellular immune response directed against the uncovered immunogenic epitopes in gp120 at the sites of the deleted glycans. In vivo studies will reveal whether this unique potential of drugs like PRM-S can be exploited in the clinical setting.
The low, if any, toxicity of PRM-S to a variety of cell lines and PBMC has been attested to by a virtual lack of influence on the induction of a variety of cytokines, chemokines, and differentiation markers by PBMC, a phenomenon that strikingly differs from the situation with the (α-1,2)mannose-specific cyanovirin, which abundantly stimulates a variety of differentiation markers and cytokines/chemokines upon exposure to PBMC cultures (6, 19). In this respect, it should also be noticed that it has been earlier demonstrated that PRM-S was not toxic in mice treated intravenously with drug doses up to 150 mg/kg of body weight (26). Also, there were no signs of toxicity of PRM-S when applied (100 μg) on the chorioallantoic membrane of fertilized chicken eggs (data not shown).
It is currently unclear how PRM-S and PRM-A exactly bind to HIV gp120. The kinetic SPR-based binding experiments revealed complex kinetics, likely due to multiple pradimicin molecules binding to one (monomeric) gp120 molecule bound on the sensor chip. Indeed, the drug off-rates (kd) clearly show biphasic kinetics starting with a fast (α) off-rate but ending with a slow (β) dissociation phase. Since the β drug dissociation phase is probably more important in terms of eventual antiviral efficacy than the α drug dissociation phase, it is of interest to note that the apparent KD values for the pradimicins were in the high nanomolar range, and PRM-S and PRM-A behaved strikingly similarly in terms of their affinities to gp120. As the studied interactions are not perfect 1:1 Langmuir, it is not possible to calculate the number of drug molecules binding to gp120. In this respect, the antibiotics differ from any other anti-HIV drug directed against other targets, such as reverse transcriptase, protease, gp41, integrase, or the cellular (co)receptors, where a single drug molecule stoichiometrically binds to a single target molecule. This phenomenon, which is so far unique for carbohydrate-binding agents such as the pradimicins, may also explain the relatively high genetic barrier of these compounds against a broad variety of HIV clades. It is interesting to note that the pradimicins also significantly interact with the transmembrane glycoprotein gp41 of HIV-1, albeit at 20- to 35-fold higher apparent KD values. Since no N-glycan deletions have ever been observed in gp41 of PRM-A-resistant (8) and PRM-S-resistant (this study) virus strains, this binding to gp41 may most likely not be directly related to the eventual antiviral activity of the pradimicins.
The pradimicins have been suggested to be physiologically active as a (dimeric) complex with Ca2+ (17, 29, 30). Experimental evidence has indeed been provided that in the presence of PRM-A, (α-1,2)mannose oligomers, and Ca2+, a ternary complex is formed that consists of one Ca2+ ion, two pradimicin-A molecules and four (α-1,2)mannose oligomers (17, 29, 30). In the absence of Ca2+, PRM-A and PRM-S are unable to bind to HIV-1 gp120 in SPR experiments. Also, methylation of the free carboxylic acid of PRM-A has been demonstrated to result in complete inactivity of the antibiotic (29), suggesting that Ca2+ may act as a bridging ion between the carboxylic acid groups of two PRM-A molecules. It is currently unclear whether Ca2+ solely plays a role in bridging two pradimicin molecules or whether it also plays a role in coordinating mannose binding to the antibiotics, as C-type lectins seem to do. These findings also suggest that at least two (α-1,2)mannose oligomer binding sites must be present at a defined distance in the pradimicin complex to allow concomitant multiple glycan binding of the antibiotics. Such a “cross-linking” capacity of glycans may be a prerequisite for the eventual antiviral action of the pradimicins. This requirement of cross-linking to afford biological activity has also been observed and reported for other lectins, such as cyanovirin N (23).
In conclusion, PRM-S consistently suppresses HIV-1 strains that belong to different viral clades, HIV-2, and SIV. It binds to the glycans of HIV-1 gp120 with a high level of affinity and prevents cell-cell fusion, virus capture by DC-SIGN-expressing cells, and subsequent HIV transmission to, and infection of, CD4+ T lymphocytes. Its characteristics of a HIV fusion inhibitor with a high genetic barrier and its high solubility that allow it to reach high local drug concentrations (i.e., intravaginally) or drug plasma levels may give it a potential therapeutic edge over other anti-HIV drug leads, in particular for microbicidal application.
This work was supported by the EMPRO no. 503558 of the 6th Frame Work Programme and CHAARM no. 242135 of the 7th Frame Work Programme of the European Commission, the Fonds voor Wetenschappelijk Onderzoek (FWO) Krediet nr. G-485-08, the Centers of Excellence of the Katholieke Universiteit at Leuven (Krediet nr. EF/05/015), and the Fondation Dormeur.
We are grateful to Ann Absillis, Leen Ingels, Lizette van Berckelaer, Yoeri Schrooten, Sandra Claes, Rebecca Provinciael, and Eric Fonteyn for excellent technical assistance and Christiane Callebaut for dedicated editorial help.
Published ahead of print on 4 January 2010.